Key Transitions in Animal Evolution

Key Transitions In Animal Evolution

© 2011 by Taylor and Francis Group, LLC

Key Transitions In Animal Evolution Editors

Rob DeSalle American Museum of Natural History Division of Invertebrate Zoology New York USA

Bernd Schierwater ITZ, Ecology and Evolution TiHo Hannover Germany

6000 Broken Sound Parkway, NW Suite 300, Boca Raton, FL 33487 Taylor & Francis Group 270 Madison Avenue New York, NY 10016 an informa business 2 Park Square, Milton Park www.crcpress.com Abingdon, Oxon OX 14 4RN, UK

CRC Press

© 2011 by Taylor and Francis Group, LLC

Science Publishers Enfield, New Hampshire

Published by Science Publishers, P.O. Box 699, Enfield, NH 03748, USA An imprint of Edenbridge Ltd., British Channel Islands E-mail: [email protected]

Website: www.scipub.net

Marketed and distributed by: 6000 Broken Sound Parkway, NW Suite 300, Boca Raton, FL 33487 Taylor & Francis Group 270 Madison Avenue New York, NY 10016 an informa business 2 Park Square, Milton Park www.crcpress.com Abingdon, Oxon OX 14 4RN, UK

CRC Press

Copyright reserved © 2011 ISBN 978-1-57808-695-5 Library of Congress Cataloging-in-Publication Data

Key transitions in animal evolution / [edited by] Rob Desalle, Bernd Schierwater. p. cm. Includes bibliographical references and index. ISBN 978-1-57808-695-5 (hardback) 1. Phylogeny. 2. Evolution (Biology) 3. Genetic transcription. I. DeSalle, Rob II. Schierwater, B. (Bernd), 1958QH367.5.K49 2010 591.3’8--dc22 2010033462

The views expressed in this book are those of the author(s) and the publisher does not assume responsibility for the authenticity of the findings/conclusions drawn by the author(s). Also no responsibility is assumed by the publishers for any damage to the property or persons as a result of operation or use of this publication and/or the information contained herein. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying or otherwise, without the prior permission of the publisher, in writing. The exception to this is when a reasonable part of the text is quoted for purpose of book review, abstracting etc. This book is sold subject to the condition that it shall not, by way of trade or otherwise be lent, re-sold, hired out, or otherwise circulated without the publisher’s prior consent in any form of binding or cover other than that in which it is published and without a similar condition including this condition being imposed on the subsequent purchaser. Printed in the United States of America

© 2011 by Taylor and Francis Group, LLC

Introduction A Phylogenomic Journey Through the Animal Tree of Life: Key Innovations in the Evolution of Metazoa Bernd Schierwater 1,2 and Rob DeSalle 2

“All animals are equal, but some animals are more equal than others.” —George Orwell “You can observe a lot just by watchin’.” —Yogi Berra The origin of multicellular animals is one of those difficult and delicate biological problems that makes careers and produces enemies. Many ideas have been proffered and many hypotheses have been tested with respect to the subject. In order to address this subject further and to assess progress in the examination of animal origins and transitions, an international group of scientists was convened at the Society for Comparative Biology meeting in 2007 in Phoenix, Arizona. Since that meeting the organizers have worked to put together a volume of essays stemming from the talks at the meetings. While the meeting occurred a few years back, we have updated and expanded all of the papers in the symposium and have added several chapters that did not occur in the original symposium. This book seeks to bring together experts in many areas of research on the origin of multicellular animals from a variety of biological perspectives. An extra dimension of this book is the addition of chapter ending questions for 1 ITZ, Ecology & Evolution, TiHo Hannover, Bünteweg 17d, D-30559 Hannover, Germany. E-mail: [email protected] 2 American Museum of Natural History, New York, Division of Invertebrate Zoology 79 St. at Central Park West, New York, NY 10024, USA. E-mail: [email protected]

© 2011 by Taylor and Francis Group, LLC

vi

Key Transitions in Animal Evolution

discussion that we hope students reading the volume will find useful in strengthening their knowledge of the individual subjects covered by the chapters. The book is divided into three sections. The first section entitled “Tangled Roots in the Animal Tree of Life” examines the phylogenetic relationships of animals from the single celled close outgroups of animals to fine tuned arthropod phylogeny. The second section entitled “The Earliest Animals: from Genes to Transitions” takes groups of living animals that represent the earliest body plans for animals and examines their evolution and development. The final section of the book entitled “Pattern and Process at the Base of the Metazoan Tree of Life” examines the life history of basal metazoans and the major transitions such as feeding, germ line evolution and adaptation to the environment, these organisms experienced in evolution. Animals and bauplans: The background on body plans for Metazoa is interesting. As DeSalle and Schierwater (2007) describe: Multicellular animals (metazoans) almost certainly derive from a cum grano salis “single” celled protozoan ancestor. Metazoa—by definition— harbor more than one somatic cell type and can be classified into basal (or diploblastic) and derived (or triploblastic) groups. These groups differ by the existence of three (triploblasts) or two germ layers only (diploblasts). It is generally accepted that triploblasts derive from a diploblastic ancestor, implying that multicellular animals evolved from simple to more complex (anagenetic evolution). The basal (diploblastic) metazoans consist of sponges, placozoans, cnidarians, and ctenophores. The higher (derived triploblastic) metazoan groups are represented by two lines which differ by the fate of the blastoporus. In the Deuterostomia lineage the blastoporus becomes the anus of the adult, in the Protostomia the blastoporus becomes the mouth of the adult. Because of the definition of ventral and dorsal according to the position of the final mouth, Protostomia and Deuterostomia represent opposite dorsal-ventral bauplans and endo- versus exogenuous skeleton systems. The above transitions from single celled to multicellular, from diploblastic to triploblastic, and from protostomic to deuterostomic are—despite substantial efforts—not yet understood. While the anatomical transitions between groups is well known, the jury on the phylogenetic relationships of the major groups of animals at the base of the metazoan tree of life is still out. Within just 12 months three papers appeared in the literature with three rather different hypotheses concerning the topology of the tree of life in this area. Dunn et al. (2008) using EST data on over 70 taxa, suggested that Ctenophores were the most basal animal, with Bilateria sister to sponges AND Cnidaria. Schierwater et al. (2009) followed with a detailed analysis of anatomy, molecules and molecular structure to come to a similar conclusion of Dunn et al. (2008)

© 2011 by Taylor and Francis Group, LLC

Introduction

vii

that Bilateria are sister to Cnidaria AND Porifera. However, by adding the Placozoa to their analysis, Schierwater et al., hypothesized that there was an early Diploblast (Placozoa, Cnidaria, Ctenophora and Porifera)— Triploblast (Bilateria) split, with Placozoa as the most basal diploblast (cf. Dellaporta et al. 2006). The final study in this trio, came from Philippe et al. (2009) who suggest that the tree of life in this area is more like traditional understanding, where Porifera are the most basal metazoan, followed by Placozoa, and then Cnidaria and Bilateria. A clue to the diversity of hypotheses for the topology in this part of the tree of life can be found in Siddall (2009) who reanalyzed all three of these data sets. Siddall points out which relationships in the three studies are robust and which are not and it is clear that much more work needs to be done on this problem. To set the stage for what the student might need to know about metazoan phylogenetics we present in this volume, several papers that discuss the metazoan part of the tree of life. Vazquez et al. start off this first section of the book entitled “Tangled Roots in the Animal Tree of Life”, with a description of phylogeny of single celled eukaryotes and how these relate to the relationships of metazoa. The outgroup status for the Metazoa is followed by two chapters that address the relationships of the major animal groups; specifically the chapter by Kocot et al. focuses on broad based nuclear gene studies and the chapter by Lavrov utilizes mtDNA whole genomes to examine the relationships of the major animal groups. To demonstrate the flexibility of phylogenetic analysis in the Metazoa, we also include a summary of the relationships of arthropod taxa by Deutsch. Comparative biology of early metazoans: The rapidly increasing number of whole genome projects has increased the number of model systems for evolutionary developmental research. As DeSalle and Schierwater (2007) suggest: Genomic features require a comparative framework to be understood, and thus the choice of the right model organisms becomes particularly crucial (Martindale 2005). Genes require a comparative classification and when animal genomes are sequenced, homologs need be assigned to various classes with some certainty (Copley et al. 2004). The interpretation of “gene homologs” has been a hot subject of debate and different researchers use crucial terms like “anterior/posterior and oral/aboral in different, and interpretive ways (e.g. Finnerty 2003, Abouheif 1999), leading to conflicting views sometimes on the same (or similar) data set (Kamm et al. 2006, Chourout et al. 2006, Ryan et al. 2007). Great expectations derived from the study of Antp-class genes to unravel the evolution of axis formation, and thus the basic bauplan setup for all major metazoan groups (e.g. Deutsch & Mouchel-Vielh 2003).

© 2011 by Taylor and Francis Group, LLC

viii

Key Transitions in Animal Evolution

Since the initial application of in situ techniques and antibody staining approaches to evolutionary questions in the 1990’s, a proliferation of studies has appeared in the literature. These studies opened the way for a better understanding of the genes involved in specific morphologies of plants and animals. The combination of genome level gene content analysis and where genes are expressed in animals have led to a real transition in how scientists look at key transitions in animal evolution. The first two chapters in this section entitled “The Earliest Animals: from Genes to Transitions”, look closely at the evolution of the nervous system. Nickel examines the biology and genomics of sponges to develop a framework for a pre-nervous system in animals. Galliot et al. examine neural genes and neurogenesis in cnidaria. Boero and Piraino use the existing ideas about development to hypothesize a one step transition from cnidaria to bilateria. This interesting hypothesis is an important example of transitional thinking. The next chapter by Jacobs et al. looks at the evolution of sensory structures and mechanisms in basal animals from an evo-devo perspective. While the nervous system and sensory apparatus are two excellent examples of transitional structure in animal evolution, the basic body plan change to bilateral symmetry has also been a major target for evo-devo research. The chapter by Ball et al. examines many of the developmental aspects of the change to a bilateral body plan. Finally, developmental mechanisms control other more specific anatomical aspects of animals. To cover all of these would be an immense undertaking so we give the student a “taste” of the kinds of studies that can be accomplished with genome level information and evo-devo approaches. Oakely and Plachetzki focus on phototransduction and how this important transition evolved in metazoa. Hox genes have been the “poster genes” for evo-devo since its inception. In order to represent the far reaching work in the area of Hox gene evolution for this volume, Kappen presents a summary of the role of Hox genes in the reproductive biology of mammals. While the subject matter of this section is vast, we hope we give the student a good background and a broad view of the kind of research being conducted. Major transitional problems at the origin of animals: Scientists can focus on specific aspects of development. Their relevance to how animal live evolved in very direct fashion as a result of genomics and the developmental approaches available. However, larger more quantitative transitions such as feeding and whole scale changes in developmental strategies such as evolution of the germ line, are also a fertile area of examination for the base of the animal tree of life. In addition, the transition from a very simple metazoan form to more complex forms can best be understood after a strong background for the simple animals have been presented. The first two papers in this section entitled “Pattern and Process at the Base of the Metazoan Tree of Life”, address the life history and biology of Placozoa.

© 2011 by Taylor and Francis Group, LLC

Introduction

ix

These chapters by Pearse and Voigt and Schierwater et al. should clearly be the reference point for understanding the biology of this little understood animal and should be an important stepping off point for anyone interested in animal evolution regardless of the phylogenetic placement of this enigmatic animal. Because the feeding apparatus of animals vary greatly across phylogeny, unraveling the events that have occurred is an important and compelling subject. Hence, Blackstone examines the evolution of feeding in animals. An understanding of how animals evolved is also dependent on the environmental conditions under which they evolved so the interrelationship of animal evolution with biogeochemical events is examined in the next chapter by Gaidos. Extavour finishes the section by examining the key events and transitions involved in the evolution of germ lines. Conclusion. ‘Watchin’: Understanding the changes that occurred at the base of the metazoan branch of the tree of life is as Yogi Bera, the famous American baseball player and philosopher once said, all about ‘watchin’; but “watchin” what? Readers of this volume will be directed to “watch” a lot of things and we hope that this book on Key Transitions in Animal Evolution contributes to the observational richness on the metazoan branch of the tree of life. It is clear that many of the questions relevant to this part of the tree of life have not been answered and some have not even been addressed. However, the more we observe (watchin’) about this question, the richer our understanding becomes and more importantly, the more well defined our questions and hypotheses become. Acknowledgments: We thank all speakers and particularly all authors for their intriguing contributions. The symposium was supported by a grant to BS and RD from the NSF (DEB-0706893) and general support from the SICB. References Abouheif, E. 1999. Establishing homology criteria for regulatory gene networks: prospects and challenges. Novartis Found. Symp. 222: 207–225. Chourrout, D. and F. Delsuc, P. Chourrout, R.B. Edvardsen, F. Rentzsch, E. Renfer, M.F. Jensen, B. Zhu, P. de Jong, R.E. Steele, and U. Technau. 2006. Minimal ProtoHox cluster inferred from bilaterian and cnidarian Hox complements. Nature 442: 684–672. Copley, R.R. and P. Aloy, R.B. Russell, and M.J. Telford. 2004. Systematic searches for molecular synapomorphies in model metazoan genomes give some support for Ecdysozoa after accounting for the idiosyncrasies of Caenorhabditis elegans. Evol. Dev. 6: 164–169. Dellaporta, S.L, and A. Xu, S. Sagasser, W. Jakob, M.A. Moreno, L.W. Buss, and B. Schierwater. 2006. Mitochondrial genome of Trichoplax adhaerens supports Placozoa as the basal lower metazoan phylum. Proc. Natl. Acad. Sci. USA 103: 8751–8756. DeSalle, R. and B. Schierwater .2007. Key transitions in animal evolution. Integr. Comp. Biol. 47: 667–669.

© 2011 by Taylor and Francis Group, LLC

x

Key Transitions in Animal Evolution

Deutsch, J.S. and E. Mouchel-Vielh. 2003. Hox genes and the crustacean body plan. Bioessays 25: 878–987. Dunn, C.W. and A. Hejnol, D.Q. Matus, K. Pang, W.E. Browne, S.A. Smith, E. Seaver, G.W. Rouse, M. Obst, G.D. Edgecombe, M.V. Sørensen, S.H. Haddock, A. Schmidt-Rhaesa A. Okusu, R.M. Kristensen, W.C. Wheeler, M.Q. Martindale, and G. Giribet. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452:745–749. Finnerty, J.R. 2003. The origins of axial patterning in the metazoa: how old is bilateral symmetry? Int. J. Dev. Biol. 47: 523–529. Kamm. K. and B. Schierwater, W. Jakob, and D. Miller. 2006. Axial patterning and diversification in the Cnidaria predate the Hox system. Curr. Biol. 16: 920–926. Martindale, M.Q. 2005. The evolution of metazoan axial properties. Nat. Rev. Genet. 6: 917–927. Philippe, H. and R. Derelle, P. Lopez, K. Pick, C. Borchiellini, N. Boury-Esnault, J. Vacelet, E. Renard, E. Houliston, E. Quéinnec, C. Da Silva, P. Wincker, H. Le Guyader, S. Leys, D.J. Jackson, F. Schreiber, D. Erpenbeck, B. Morgenstern, G. Wörheide, and M. Manuel. 2009. Phylogenomics Revives Traditional Views on Deep Animal Relationships. Curr. Biol. 19: 706–712. Ryan, J.F. and M.E. Mazza, K. Pang, D.Q. Matus, A.D. Baxevanis, M.Q. Martindale, and J.R. Finnerty. 2007. Pre-bilaterian origins of the Hox cluster and the Hox code: evidence from the sea anemone, Nematostella vectensis. PLoS ONE 2: 1–23. Schierwater, B. and M. Eitel, W. Jakob, H.-J. Osigus, H. Hadrys, S. Dellaporta, S.O. Kolokotronis, and R. DeSalle. 2009. Concatenated Molecular and Morphological Analysis Sheds Light on Early Metazoan Evolution and Fuels a Modern “Urmetazoon” Hypothesis. PlosBiology 7: e1000020. Siddall, M.E. 2009. Unringing a bell: metazoan phylogenomics and the partition bootstrap. Cladistics 25: 1–9.

© 2011 by Taylor and Francis Group, LLC

Contents Introduction. A Phylogenomic Journey Through the Animal Tree of Life: Key Innovations in the Evolution of Metazoa Bernd Schierwater and Rob DeSalle

v

Section 1. Tangled Roots in the Animal Tree of Life 1. Putting Animals in their Place Within a Context of Eukaryotic Innovations Danielle Vazquez, Laura Wegener Parfrey, and Laura A. Katz

3

2. Elucidating Animal Phylogeny: Advances in Knowledge and Forthcoming Challenges Kevin M. Kocot, Johanna T. Cannon, and Kenneth. M. Halanych

15

3. Key Transitions in Animal Evolution: a Mitochondrial DNA Perspective Dennis V. Lavrov

34

4. Pending Issues in Development and Phylogeny of Arthropods Jean S. Deutsch

54

Section 2. The Earliest Animals: From Genes to Transitions 5. The Pre-Nervous System and Beyond—Poriferan Milestones in the Early Evolution of the Metazoan Nervous System Michael Nickel

85

6. A Key Innovation in Animal Evolution, the Emergence of Neurogenesis: Cellular and Molecular Cues from Cnidarian Nervous Systems Brigitte Galliot

127

7. From Cnidaria to “Higher Metazoa” in One Step Ferdinando Boero and Stefano Piraino

162

© 2011 by Taylor and Francis Group, LLC

xii

Key Transitions in Animal Evolution

8. Basal Metazoan Sensory Evolution D.K. Jacobs, D.A. Gold, N. Nakanishi, D. Yuan, A. Camara, S.A. Nichols, and V. Hartenstein

175

9. Cnidarian Gene Expression Patterns and the Origins of 197 Bilaterality—Are Cnidarians Reading the Same Game Plan as “Higher” Animals? Eldon E. Ball, Danielle M. de Jong, Bernd Schierwater, Chuya Shinzato, David C. Hayward and David J. Miller 10. Key Transitions During Animal Phototransduction Evolution: Co-duplication as a Null Hypothesis for the Evolutionary Origins of Novel Traits Todd H. Oakley and David C. Plachetzki

217

11. Vertebrate Hox Genes and Specializations in Mammals Claudia Kappen

238

Section 3. Pattern and Process at the Base of the Metazoan Tree of Life 12. Field Biology of Placozoans (Trichoplax): Distribution, Diversity, Biotic Interactions Vicki Buchsbaum Pearse and Oliver Voigt

259

13. Trichoplax and Placozoa: One of the Crucial Keys to Understanding Metazoan Evolution

289

Bernd Schierwater, Michael Eitel, Hans-Jürgen Osigus, Karolin von der Chevallerie, Tjard Bergmann, Heike Hadrys, Maria Cramm, Laura Heck, Wolfgang Jakob, Michael R. Lang, Rob DeSalle 14. A Food’s-Eye View of Animal Transitions Neil W. Blackstone

327

15. Lost in Transition: The Biogeochemical Context of Animal Origins Eric Gaidos

345

16. Redefining Stem Cells and Assembling Germ Plasm: Key Transitions in the Evolution of the Germ Line John Srouji and Cassandra Extavour

360

17. Questions and Discussion Index Color Plate Section

398 409 417

© 2011 by Taylor and Francis Group, LLC

Section 1 Tangled Roots in the Animal Tree of Life

© 2011 by Taylor and Francis Group, LLC

Chapter 1

Putting Animals in their Place within a Context of Eukaryotic Innovations Danielle Vazquez,1 Laura Wegener Parfrey 2 and Laura A. Katz 1,2

Animals represent just one of an estimated 70+ lineages of eukaryotes (Patterson 1999), and the bulk of the remaining lineages are microbial. During the past decade, perspectives of the organization of eukaryotic diversity have undergone a major shift to a system that recognizes five or six high-level groupings (Adl et al. 2005, Tekle et al. 2009). While deep nodes on the eukaryotic tree of life remain elusive (reviewed in Parfrey et al. 2006), the position of animals emerging from within microbial lineages has been robustly resolved by several lines of evidence (King 2004). Animals are placed within the supergroup Opisthokonta, along with fungi and numerous microbial relatives (Adl et al. 2005, Steenkamp et al. 2006). In this chapter we place animals in their phylogenetic context by focusing on lineages of eukaryotes defined by one or more ultrastructural identities (i.e., unique combinations of subcellular structures). Many of these clades are marked by major innovations in cell and/or genome structure at their base. We argue that this contrasts with animals, which lack clear innovations at the base of the lineage despite many within (e.g., triploblasty, true gut). We focus on eight exemplar clades: foraminifera, diatoms, ciliates, apicomplexa, dinoflagellates, euglenids, parabasalids

1

Department of Biological Sciences, Smith College, Northampton, MA, USA. Program in Organismic and Evolutionary Biology, University of Massachusetts, Amherst, MA, USA. E-mail: [email protected] 2

© 2011 by Taylor and Francis Group, LLC

4

Key Transitions in Animal Evolution

and kinetoplastids. Many of the eukaryotic clades originally defined by morphological and/or ultrastructural characteristics are now grouped into larger clades—the Stramenopiles, Alveolates, and Euglenozoa—that are well-supported in molecular analyses. Here, we aim to accomplish two goals: (1) putting animals into an evolutionary context by exploring major transitions at the base of diverse microbial clades; and (2) using examples from diverse eukaryotic lineages to highlight the complex evolutionary patterns of features that are often believed to be fixed across the eukaryotic tree or unique to animals (as well as plants and fungi). In both sections, we aim to be exemplary rather than exhaustive and our intent is to provide a broader context for interpreting major transitions within animals, the subject of the remainder of this book. Part one: Understanding Innovations Across the Eukaryotic Tree of Life We have chosen a limited set of clades to demonstrate the types of transitions that have occurred at the base of many microbial lineages. These examples were chosen partially due to the availability of data and partially based on the authors’ areas of expertise. Diatoms are among the most diverse clades of eukaryotes, with an estimated 100,000 extant species, and fall within the stramenopiles (Fig. 1). The stramenopiles are a well-supported clade that includes kelp (Phaeophyceae), watermolds (oomycetes), slime nets (labyrinthulids), and many other lineages (Andersen 2004). Synapomorphies for diatoms include the presence of a silica shell, which gives diatoms their distinctive appearance (Fig. 2a). These shells are thought to have evolved initially as a byproduct of silica metabolism, and appear to play a protective role (Schmid 2003). Three of our exemplar lineages—ciliates, dinoflagellates, and apicomplexa—are members of a single clade, the alveolates (Fig. 1). Alveolates are united by the presence of alveoli, a system of abutting sacs that form a layer immediately underneath the plasma membrane that is supported by microtubules (Patterson 1999). These alveolar sacs are thought to add rigidity to the cells. Ciliates (Fig. 2b) are marked by numerous innovations. Morphologically, all ciliates contain rows of cilia (flagella) on the cell surface that are supported by a complex series of microtubules (Lynn 1996). The diverse patterns of ciliary organization have given this clade tremendous morphological complexity. The organization of cilia also provides numerous characters by which ciliates are described, classified, and monographed (e.g. Corliss

© 2011 by Taylor and Francis Group, LLC

Putting Animals in their Place within a Context of Eukaryotic Innovations

Foraminifera

5

Variant Mitosis Multicellular + Rhizaria

Cercozoa

+

Brown Algae

+

Diatoms

+

Apicomplexa

+

Dinoflagellates

+

Ciliates

+

+

Green Algae

+

+

Euglenozoa

+

Parabasalia

+

Entamoeba

+

Eumycetozoa

+

+

Fungi

+

+

Microsporidia

+

Nucleariids

+

+ Stramenopiles

Alveolates

Excavata

Amoebozoa

Opisthokonta

Choanoflagellates Animals

+

Fig. 1. Distribution of multicellularity and alternative types of mitosis across the eukaryotic tree of life. Variant mitosis: + denotes lineages in which mitosis deviates from the open orthomitosis of at least some members (Fig. 3), the “classic” animal and plant model. Multicellar: Clades with one or more members that have multicellular life cycle stages. Tree modified from Tekle et al. 2009.

1979). Ciliates are also defined by the presence of two distinct types of nuclei within each cell—the germline micronucleus and the somatic macronucleus. The genomic implications of this feature are discussed below. Apicomplexa, a second major lineage within the Alveolates, include the malarial parasites Plasmodium falciparum (Fig. 2c) and P. vivax. Apicomplexa are defined by the presence of an ‘apical complex’ at the anterior end of

© 2011 by Taylor and Francis Group, LLC

6

Key Transitions in Animal Evolution

the cells (Morrison 2009). This structure consists of a ring of specialized microtubules and an underlying system of modified vesicles, which secrete chemicals to break down host cell walls. The apical complex enables these parasites to invade a variety of tissues and cells, contributing to the success of this diverse clade (Morrison 2009). Dinoflagellates (Fig. 2d), the third major alveolate lineage, are marked by innovations including a very unusual nucleus, called a dinokaryon. In most eukaryotes, nuclear DNA wraps around histone proteins, forming a nucleosome that can be further packaged depending on the degree of chromosome condensation. Most nuclei have condensed (heterochromatin) and decondensed (euchromatin) chromosome regions, the proportions of which vary according to transcriptional activity and life cycle stage. In contrast, the dinoflagellate nuclei are devoid of canonical histones— though histone-like proteins have been identified (Wong et al. 2003)—and their chromosomes are permanently condensed (Hackett et al. 2004). The dinoflagellate chromosome structure is proposed to be a cholesteric liquid crystal (Rizzo 2003, Costas and Goyanes 2005), but the implications of these features are not known. Dinoflagellate nuclei divide by closed mitosis (see below), with the ability to create tunnels for microtubules (Hackett et al.2004). Foraminifera (Fig. 1, Fig. 2e) are defined by their granular reticulopodia (Bowser and Travis 2002), a dynamic network of branching and anastomosing pseudopodia that supports bidirectional movement of particles both within and on the surface of the pseudopodia. Reticulopodia are supported by an infrastructure of microtubule bundles that enable the dynamic motility of the network (Bowser and Travis 2002). The reticulopodia and attached particles move several orders of magnitude faster than the pseudopods of other amoeboid organisms. Unique microtubule assembly and disassembly underlain by divergent tubulins is hypothesized to enable such speed (Habura et al. 2005). Ultrastructural studies of the reticulopodia show that microtubules disassemble into helical filaments, unique tubulin polymorphs that can be stored as paracrystals and shipped out to rapidly growing points of the network (Bowser and Travis 2002). Foraminifera interact with their environment through the reticulopodia by capturing food, building shells (also called tests), and moving. The innovation of the reticulopodia is thought to have enabled the diversification and ecological success within the clade. The supergroup Excavata (Fig. 1), as proposed by Cavalier-Smith (2002), seeks to unite a number of protist lineages, including the euglenids, kinetoplastids, and parabasalids. All members are either excavate taxa (they possess an excavated ventral feeding groove) or are thought to be

© 2011 by Taylor and Francis Group, LLC

Putting Animals in their Place within a Context of Eukaryotic Innovations

7

descended from excavates, though the monophyly of the supergroup remains disputed (Simpson 2003). We describe a few of these lineages below. Together, the kinetoplastids and euglenids comprise the bulk of the Euglenozoa, a taxon whose monophyly is robust (Simpson 2003, Parfrey et al. 2006). The kinetoplastids include marine flagellates such as Rhynchomonas nasuta (Fig. 2f) as well as the trypanosomes, parasites responsible for serious human illnesses such as sleeping sickness (Trypanosoma brucei) and leishmaniasis (e.g. Leishmania major) (Simpson et al. 2002). They are distinguished from other protist lineages by the presence of a single mitochondrion and a unique organization of the mitochondrial genome. Kinetoplastids have a condensed body of mitochondrial DNA called the kinetoplast that is large enough to be detected by light microscopy and is the source of group’s name (Simpson et al. 2002). In trypanosomes, the kinetoplast DNA (kDNA) is arranged into interlocking maxi- and minicircles. Maxicircles are few in number and encode incomplete copies of typical mitochondrial genes, whereas the minicircles can number in the thousands (Simpson et al. 2002). Each minicircle encodes 1 to 4 small guide RNAs that mediate the extensive editing of maxicircle transcripts. Also gene organization along the nuclear chromosomes of kinetoplastids follows an unusual pattern termed absolute strand polarity, in which the genes are arranged in large clusters along just one strand of the DNA helix (McGrath and Katz 2004). In contrast, the genes of most other organisms are encoded along both strands. The euglenids (Fig. 2g) are mostly free-living freshwater flagellates characterized by a unique pellicle made up of proteinaceous strips located beneath the cell membrane (Leedale 1967). The pellicle is further supported by a network of microtubules (Leedale 1967). While some move only using their flagella, in many euglenids the pellicle strips can slip past each other, thereby producing a distinctive inching motion called metaboly. Parabasalids are a flagellate lineage composed of roughly 80 amitochondriate genera (Brugerolle and Müller 2000) such as the well-studied Trichomonas vaginalis and Tritrichomonas muris (Fig. 2h). Most members are either parasites or endosymbionts of other animals, notably termites and humans. The taxon is distinguished by their prominent Golgi bodies, which are linked to the basal bodies by parabasal fibers (Brugerolle and Müller 2000). Animals are members of the Opisthokonta, a robust clade defined by both a morphological and molecular synapomorphy. The morphological synapomorphy is the presence of a single posterior flagellum, which is found in animal sperm cells as well as choanoflagellates, the closest relatives of animals. The similarity of cell architecture and flagellar

© 2011 by Taylor and Francis Group, LLC

8

Key Transitions in Animal Evolution

position between sponges and choanoflagellates was first noticed more than a century ago (King 2004). This character has been lost in some Opisthokont groups including the fungi. A molecular synapomorphy for this group is the presence of an insertion in the Elongation factor 1a (EF1a) gene (Baldauf and Palmer 1993), for those taxa that have the canonical EF1a (Keeling and Inagaki 2004, Gile et al. 2006).

Fig. 2. Representative taxa from major eukaryotic lineages discussed. Stramenopiles: (a) Triceratium pentacrinus, a diatom; Alveolates (b-d): (b) Trithigmostoma cucullulus, a ciliate, (c) Plasmodium falciparum, an apicomplexan, and (d) Akashiwo sanguinea, a dinoflagellate; (e) Ammonia sp., a foraminiferan; Excavata: (f) Rhynchomonas nasuta, a kinetoplastid and (g) Euglena mutabilis, a euglenid; (h) Tritrichomonas muris, a parabasalid; Opisthokonta: (i) Conochilus sp., a colonial rotifer. Scale bars: b-d,f-h = 10 micrometers; a, e = 100 micrometers; i = 200 micrometers. All images are provided by micro*scope (http: //starcentral.mbl.edu/ microscope/portal.php) except c, which is used under PLoS Biology’s Open Access rules and can be found in Lacroix et al. (2005), credit Dr. Mae Melvin. Color image of this figure appears in the color plate section at the end of the book.

© 2011 by Taylor and Francis Group, LLC

Putting Animals in their Place within a Context of Eukaryotic Innovations

9

Several lineages of eukaryotes are emerging from molecular studies that do not have morphological synapomorphies. These lineages, including Cercozoa, Amoebozoa, and Rhizaria, unite a surprising diversity of morphological forms. For example, molecular studies have placed more than 15 lineages of amoebae, parasites, flagellates, and heliozoa with ultrastructural identities into the Cercozoa. These include cercomonads (gliding flagellates), and chlorarachniophytes (filose amoebae with green algal plastids), and desmothoracids (heliozoa). The lack of a discernable morphological innovation uniting these groups may be a result of the ancient age of these clades. Part two: Microbial Lineages Defy Many of the Long-Held Assumptions About Eukaryotes Multicellularity Although multicellularity has often been thought of as the exclusive territory of plants, fungi, and animals, there are, in fact, a number of microbial eukaryotic lineages that exhibit the trait. The largest protists are found among the brown algae, with giant kelps such as Macrocystis reaching lengths of up to 60 meters (Bonner 1998). A number of brown algae also undergo extensive cell differentiation, with some producing not only stem- and leaf-like parts, but also gas-filled bladders. Furthermore, the majority of giant kelps have photosynthetic filaments at the very edges of their stalks and blades, an example of tissue differentiation. Other lineages achieve muticellularity through mechanisms that are quite different from what one finds in animals, plants, and fungi. Some of them are more colonial rather than strictly multicellular: in the ciliate genus Zoothamnium, individual cells come together to form branched colonies. The cells are held together such that should one or a few cells contract, the rest of the colony will follow suit, presumably to avoid danger (Bonner 1998). The green algae genus Volvox, on the other hand, produces striking spherical colonies made up of a collection of individual cells held together by a gelatinous glycoprotein. Colonial species are generally not considered to be truly multicellular since they do not undergo functional tissue differentiation. However, the volvocine species that form larger colonies (up to 50,000 cells) actually do experience some functional differentiation, though it is usually limited to the formation of somatic and germ cells (Herron and Michod 2008). Perhaps the most radically different approach to multicellularity is via aggregation, whereby individual cells come together (usually when conditions are poor). Unlike macrobes, though, protists that exhibit aggregative multicellularity nevertheless retain the ability to revert to their unicellular state. Dictyostelium, a genus of cellular slime mold, is one

© 2011 by Taylor and Francis Group, LLC

10

Key Transitions in Animal Evolution

of the more extensively studied of the aggregative protist lineages. These amoebae have a sexual life cycle during which they feed as independent cells, coming together into a multicellular slug only after their food supply has been depleted (Bonner 1998). Cells then collectively form a fruiting body containing environmentally resistant spores. Such aggregation is characteristic of dictyostelid amoebae as well as the acrasids (the acellular “slime molds”), but it has also been found among the ciliates. Sorogena stoianovitchae, which lives in soil and feeds on other ciliates, is a unique ciliate species in that it forms aerial fruiting bodies in the face of nutrient scarcity (Lasek-Nesselquist et al. 2001). Dynamic genomes Much of our understanding of the structure and function of genomes is derived from model animals, fungi, and plants. However, this macrobial bias overlooks the tremendous diversity of genome structure and content present in microbial eukaryotes. Data from microbial eukaryotes are challenging the assumption that genomes are largely static within species. In contrast to the textbook view, there is indeed intraspecific variation in genome architecture—the content and organization of nuclear DNA—within individuals during the life cycle and among individuals in populations (Parfrey et al. 2008). In many lineages, genome content changes during the life cycle through genome processing. Genome processing is the fragmentation, amplification, and/ or elimination of chromosomes or portions of chromosomes. Examples include: (1) ribosomal DNA amplification in numerous eukaryotic lineages (McGrath and Katz 2004, Zufall et al. 2005) and (2) whole genome rearrangements that occur in the somatic nuclei of ciliates and some animals (Prescott 1994, Jahn and Klobutcher 2002). As mentioned above, every ciliate has two types of nuclei within a single cell: a germline micronucleus and somatic macronucleus. Micronucleuei go through canonical mitosis and meiosis, but are transcriptionally silent. The transcriptionally active macronucleus develops from the micronucleus through extensive genome fragmentation and amplification. In some types of ciliates this process results in a macronucleus with ~25 million genesized chromosomes (Prescott 1994). Genome processing has also been shown to affect patterns of molecular evolution in ciliates, with rates of protein evolution correlated with the extent of chromosomal processing. In other words, ciliates with extensively processed chromosomes have more divergent proteins (Katz et al. 2004, Zufall et al. 2006). The genomes of Entamoeba vary from a ploidy of 4N to 40N in a single population (Lohia 2003). Populations can be synchronized to 4N, but the distribution of ploidy levels is restored within hours of normal growth

© 2011 by Taylor and Francis Group, LLC

Putting Animals in their Place within a Context of Eukaryotic Innovations

11

(Lohia 2003). Entamoebae also process their genomes by extensively amplifying their ribosomal DNA. They do not have any chromosomal copies of rDNA; instead, it exists only as a plasmid that varies in copy number (McGrath and Katz 2004). Foraminifera and some related lineages go through a “nuclear cleansing” process prior to reproduction that indicates genome processing occurred during the development of the nucleus. During this process, termed Zerfall, nuclear contents including DNA are condensed, ejected from the nucleus, and degraded (Føyn 1936). The remaining nuclear material divides into hundreds to thousands of gametic nuclei (Goldstein 1997). Variations in mitosis Mitosis in eukaryotes assumes a diversity of forms, in contrast to the tidy, static process familiar from textbooks. In the “classic” form of mitosis— called eumitosis or open orthomitosis—the nuclear envelope breaks down completely and chromosomes are segregated by polar spindles into two daughter cells. While open orthomitosis is characteristic of animals and plants, it actually represents only one of the many variations of mitotic division found across the eukaryotic tree of life (Fig. 3 (Heath 1980, Raikov 1982)). There are a number of ways to distinguish different types of mitosis, including variations in microtubule quantity and chromatin condensation and organization (Heath 1980) as well as nuclear pore assembly (De Souza and Osmani 2007). Nevertheless, the major forms of mitosis (Fig. 3) are defined mainly by the behavior of the nuclear envelope and the

Fig. 3. The major forms of mitosis, as defined by the degree of nuclear envelope disintegration and spindle symmetry. Redrawn from Raikov 1994.

© 2011 by Taylor and Francis Group, LLC

12

Key Transitions in Animal Evolution

symmetry of the spindles (Raikov 1994). In this section we will briefly discuss differences in both. The terms “open,” “semiopen,” and “closed” describe the degree to which the nuclear envelope has disintegrated, while “orthomitosis” and “pleuromitosis” refer to the symmetry (or lack thereof) of the mitotic spindles. Thus, in both semiopen orthomitosis and semiopen pleuromitosis, the nuclear envelope only disintegrates at the polar zones, leaving just enough space through which microtubules can enter the nucleus (Raikov 1982). In some cases, though, the nuclear envelope reseals itself soon after the microtubules have passed through (Heath 1980). Semiopen pleuromitosis differs from semiopen orthomitosis in that while the latter’s microtubules produce a bipolar, symmetrical spindle—the definition of orthomitosis —, there are two independent spindles present in pleuromitosis (Raikov 1982). The spindles start out adjacent to one another but later move away at an angle, which initiates anaphase: there is neither a metaphase nor an equatorial plate in pleuromitosis (Raikov 1982). Semiopen pleuromitosis is typical of many apicomplexans (Raikov 1982), including the malarial parasites of the genus Plasmodium. Semiopen orthomitosis, meanwhile, is typical of the green flagellates Volvocales and Chloromonadida, as well as several gregarines and heliozoans (Raikov 1982). One rare variant of “classic” open mitosis has only been observed in the gregarine genus Stylocephalus, in which the nuclear envelope completely disintegrates simply to reform around each individual chromosome (Heath 1980). All of the other major types of mitosis are closed—that is, the nuclear envelope does not disintegrate at all. In both closed intranuclear pleuromitosis and orthomitosis, the spindle remains entirely within the nucleus, though its positioning is different (Raikov 1982). Closed intranuclear pleuromitosis is typical of kinetoplastids, oxymonadids, foraminifera, radiolarians, as well as some green flagellates. Closed orthomitosis, on the other hand, is most typical of amoebae, ciliates, and some microsporidians (Raikov 1982). With extranuclear pleuromitosis, the spindle apparatus is found in the cytoplasm and contacts chromosomes through the nuclear envelope (Heath 1980, Ribeiro et al. 2000). Extranuclear pleuromitosis is typical of the parabasalids, such as the trichomonads (Raikov 1982) and dinoflagellates. Summary Here we have placed the evolutionary transitions leading to animals in the context of their microbial relatives on the eukaryotic tree of life. Microbial lineages make up the breadth of eukaryotic diversity, and many are marked by innovations in cell and/or genome structure at their base, which we believe contrasts with the origin of animals. An understanding of

© 2011 by Taylor and Francis Group, LLC

Putting Animals in their Place within a Context of Eukaryotic Innovations

13

the diversity of microbial eukaryotes provides perspective in interpreting the evolution of eukaryotic features that are typified by animals, including multicellularity, genome structure, and mitosis, as discussed here. These features highlight the variation and diversity within eukaryotes. References Adl, S.M. and A.G.B. Simpson, M.A. Farmer, R.A. Andersen, O.R. Anderson, J.R. Barta, S.S. Bowser, G. Brugerolle, R.A. Fensome, S. Fredericq, T.Y. James, S. Karpov, P. Kugrens, J. Krug, C.E. Lane, L.A. Lewis, J. Lodge, D.H. Lynn, D.G. Mann, R.M. McCourt, L. Mendoza, O. Moestrup, S.E. Mozley-Standridge, T.A. Nerad, C.A. Shearer, A.V. Smirnov, F.W. Spiegel, and M. Taylor. 2005. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J. Eukaryot. Microbiol. 52: 399–451. Andersen, R.A. 2004. Biology and systematics of heterokont and haptophyte algae. Am J. Bot. 91: 1508–1522. Baldauf, S.L. and J.D. Palmer. 1993. Animals and fungi are each other’s closest relatives: congruent evidence from multiple proteins. Proc. Natl. Acad. Sci. USA 90: 11558– 11562. Bonner, J.T. 1998. The origins of multicellularity. Integr. Biol. 1: 27–36. Bowser, S.S. and J.L. Travis. 2002a. Reticulopodia: Structural and behavioral basis for the suprageneric placement of Granuloreticulosan protists. J. Foram Res. 32: 440–447. Brugerolle, G. and M. Müller. Amitochondriate flagellates. pp. 166–189. In: J.C. Green and B.S.C. Leadbeater. [eds.] 2000. Flagellates: Unity, Diversity and Evolution. Taylor and Francis, London. Cavalier-Smith, T. 2002. The phagotrophic origin of eukaryotes and phylogenetic classification of Protozoa. Int. J. Syst. Evol. Microbiol. 52a: 297–354. Corliss, J.O. 1979. The Ciliated Protozoa: Characterization, Classification and Guide to the Literature. Pergamon Press, Oxford. Costas, E. and V. Goyanes. 2005. Architecture and evolution of dinoflagellate chromosomes: an enigmatic origin. Cytogenet. Genome Res. 109: 268–275. De Souza, C.P.C. and S.A. Osmani. 2007. Mitosis, not just open or closed. Eukaryot Cell 6: 1521–1527. Føyn, B. 1936. Über die Kernverhaltnisse der Foraminifere Myxotheca arenilega Schaudinn. Arch. Protistenkd 87: 272–295. Gile, G.H. and N.J. Patron, and P.J. Keeling. 2006. EFL GTPase in cryptomonads and the distribution of EFL and EF-1alpha in chromalveolates. Protist. 157: 435–444. Goldstein, S.T. 1997. Gametogenesis and the antiquity of reproductive pattern in the Foraminiferida. J. Foram Res. 27: 319–328. Habura, A. and L. Wegener, J.L. Travis, and S.S. Bowser. 2005. Structural and functional implications of an unusual Foraminiferal beta-tubulin. Mol. Biol. Evol. 22: 2000–2009. Hackett, J.D. and H.S. Yoon, M.B. Soares, M.F. Bonaldo, T. Casavant, T.E. Scheetz, T. Nosenko, and D. Bhattacharya. 2004. Migration of the plastid genome to the nucleus in a pridinin dinoflagellate. Curr. Biol. 14: 213–218. Heath, B. 1980. Variant mitosis in lower eukaryotes: indicators of the evolution of mitosis? Int. Rev. Cytol. 64: 1–80. Herron, M.D. and R.E. Michod. 2008. Evolution of complexity in the volvocine algae: Transitions in individuality through Darwin’s eye. Evolution 62: 436–451. Jahn, C.L. and L.A. Klobutcher. 2002. Genome remodeling in ciliated protozoa. Annu Rev Microbiol 56: 489–520. Katz, L.A. and J.G. Bornstein, E. Lasek-Nesselquist, and S.V. Muse. 2004. Dramatic diversity of ciliate histone H4 genes revealed by comparisons of patterns of substitutions and paralog divergences among eukaryotes. Mol. Biol. Evol. 21: 555–562.

© 2011 by Taylor and Francis Group, LLC

14

Key Transitions in Animal Evolution

Keeling, P.J. and Y. Inagaki. 2004. A class of eukaryotic GTPase with a punctate distribution suggesting multiple functional replacements of translation elongation factor-1 alpha. Proc. Natl. Acad. Sci. USA 101: 15380–15385. King, N. 2004. The unicellular ancestry of animal development. Dev. Cell 7: 313–325. Lacroix, R. and W.R. Mukabana, L.C. Gouagna, and J.C. Koella. 2005. Malaria infection increases attractiveness of humans to mosquitoes. PLoS Biol. 3: 1590–1593. Lasek-Nesselquist, E. and L.A. Katz. 2001. Phylogenetic position of Sorogena stoianovitchae and relationships within the class Colpodea (Ciliophora) based on SSU rDNA sequences. J. Eukaryot. Microbiol. 48: 604–607. Leedale, G.F. 1967. Euglenoid Flagellates. Prentice-Hall, Inc Englewood Cliffs N.J., pp. 96–107. Lohia, A. 2003. The cell cycle of Entamoeba histolytica. Mol Cell Biochem 253: 217–222. Lynn, D.H. Systematics of ciliates. In: K. Hausmann and P.C. Bradbury. [eds.] 1996. Ciliates: Cells as Organisms. Gustav Fischer Verlag, Stuttgart. McGrath, C.L. and L.A. Katz. 2004. Genome diversity in microbial eukaryotes. Trends Ecol. Evol. 19: 32–38. Morrison, D.A. 2009. Evolution of the Apicomplexa: where are we now? Trends in Parasitol 25: 375–382. Parfrey, L.W. and E. Barbero, E. Lasser, M. Dunthorn, D. Bhattacharya, D.J. Patterson, and L.A. Katz. 2006. Evaluating support for the current classification of eukaryotic diversity. PLoS Genet. 2: 2062–2073. Parfrey, L.W. and D.J.G. Lahr, and L.A. Katz. 2008. The dynamic nature of eukaryotic genomes. Mol Biol Evol 25: 787–794. Patterson, D.J. 1999. The diversity of eukaryotes. Amer. Nat. 154: S96–S124. Prescott, D.M. 1994. The DNA of ciliated protozoa. Microbiol. Rev. 58: 233–267. Raikov, I.B. 1982. The Protozoan Nucleus: Morphology and Evolution. Springer-Verlag, Wien. Raikov, I.B. 1994. The Diversity of Forms of Mitosis in Protozoa: A Comparative Review. European J. Protistol. 30: 253–269. Ribeiro, K.C. and L.H. Monteiro-Leal, and M. Benchimol. 2000. Contributions of the axostyle and flagella to closed mitosis in the protists Tritrichomonas foetus and Trichomonas vaginalis. J. Eukaryot. Microbiol. 47: 481–492. Rizzo, P. 2003. Those amazing dinoflagellate chromosomes. Cell Res 13: 215–217. Schmid, A.-M.M. 2003. The evolution of the silicified diatom cell wall—revisited. Diatom. Res. 17: 345–351. Simpson, A.G.B. and J. Lukes, and A.J. Roger. 2002. The evolutionary history of kinetoplastids and their kinetoplasts. Mol. Biol. Evol. 19: 2071–2083. Simpson, A.G.B. 2003. Cytoskeletal organization, phylogenetic affinities and systematics in the contentious taxon Excavata (Eukaryota). Int J Syst Evol Microbiol 53: 1759–1777. Steenkamp, E.T. and J. Wright, and S.L. Baldauf. 2006. The protistan origins of animals and fungi. Mol. Biol. Evol. 23: 93–106. Tekle, Y.I. and L.W. Parfrey, and L.A. Katz. 2009. Molecular data are transforming hypotheses on the origin and diversification of eukaryotes. Bioscience 59: 471–481. Wong, J.T.Y. and D.C. New, J.C.W. Wong, and V.K.L. Hung. 2003. Histone-like proteins of the dinoflagellate Crypthecodinium cohnii have homologies to bacterial DNA-binding proteins. Eukaryot. Cell 2: 646–650. Zufall, R.A. and T. Robinson, and L.A. Katz. 2005. Evolution of developmentally regulated genome rearrangements in eukaryotes. Journal of Experimental Zoology Part B-Molecular and Developmental Evolution 304B: 448–455. Zufall, R.A. and C.L. McGrath, S.V. Muse, and L.A. Katz. 2006. Genome architecture drives protein evolution in ciliates. Molecular Biology and Evolution 23: 1681–1687.

© 2011 by Taylor and Francis Group, LLC

Chapter 2

Elucidating Animal Phylogeny: Advances in Knowledge and Forthcoming Challenges Kevin M. Kocot, Johanna T. Cannon, and Kenneth. M. Halanych

Introduction Understanding key transitions in animal evolution is dependent in on an accurate evolutionary framework or phylogeny. Molecular phylogenetics greatly informed the study of animal evolutionary relationships during the late 1980s and 1990s and brought about several major changes to the animal tree of life (e.g. Lophotrochozoa, Ecdysozoa). More recently, our understanding of higher-level animal evolutionary relationships has been further resolved, although many questions remain unanswered. Major recent changes to our view of the animal tree of life include the placement of Acoela and Nemertodermatida at the base of Bilateria, the inclusion of sipunculans within Annelida, the placement of Onychophora as the sister taxon of Arthropoda, and the surprising find of a clade composed of Craniata and Tunicata to the exclusion of Cephalochordata. Despite these advances, a number of issues remain unresolved; including which of the basal metazoan phyla (Porifera, Placozoa, Cnidaria, or Ctenophora) is basal-most, the branching order of the three major lineages of Arthropoda, the relationships within Lophotrochozoa, and the placement of several socalled minor groups such as Chaetognatha. Here, we review the current understanding of animal phylogeny with emphasis on the most recent work and illustrate remaining gaps in our knowledge. Department of Biological Sciences, 101 Rouse Life Science Building, Auburn University, Auburn, AL USA 36849. E-mail: [email protected], [email protected], [email protected]

© 2011 by Taylor and Francis Group, LLC

16

Key Transitions in Animal Evolution

One of the goals of this volume is to examine major transitions in form or function that allowed animals to become more evolutionarily successful (as judged by either number of species, diversity, or dominance). If such transitions occur in stem or ancestral lineages and are present in all or most of the descendents, we typically call them major, or key, transitions. Understanding of such transitions is dependent upon an accurate assessment of evolutionary history and in particular, the phylogenetic relationships of the organisms being examined. Without an accurate phylogenetic framework, we have little hope of assessing a number of evolutionary issues relating to such transitions, for example: Which is the most derived taxon that shows the pre-transition state? Was the transition a single major event or a graded series of events? Has the transition in question occurred once or multiple times? Does the transition represent a de novo evolutionary invention or just a rearrangement of parts? The goal of this chapter is to address the current understanding of animal phylogeny so that key transitions can be placed in an appropriate evolutionary context. Although our understanding of animal phylogeny is far from complete, the past several years have seen significant advances in resolving animal phylogeny at all hierarchical levels. The advent of molecular approaches in the late 1980s to 1990s radically reshaped traditional hypotheses of animal phylogeny (e.g. Hyman 1959, Willmer 1990, Nielsen 2001). Several reviews (e.g. Halanych 1999, Eernisse and Peterson 2004, Halanych 2004, Giribet et al. 2007) have outlined advances based on molecular data. Halanych (2004) provided a summary of major findings from phylogenetic studies up until that time, most of which were molecular studies based on one to three genes. Due to space limitations, this contribution will essentially continue from 2004. Since then a number of multiple-gene studies have been conducted which have provided considerable insight into animal phylogenetics while raising new questions. Herein, we will consider some advantages and disadvantages of studies utilizing genome or transcriptome-level data (a.k.a. phylogenomics), provide an overview of the current understanding of metazoan phylogenetics, and discuss unresolved issues. Molecular Approaches Several of the major changes in our understanding of animal phylogeny in the late twentieth century were initially hypothesized on the basis of the nuclear ribosomal small subunit (SSU or 18S) rDNA gene (e.g. Field et al., 1988, Halanych et al. 1995, Aquinaldo et al. 1997, Ruiz-Trillo et al. 1999). The advantages of this gene include the presence of both variable and conserved regions, the latter of which facilitate oligonucleotide primer design for polymerase chain reaction (PCR). This marker also has some

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

17

potential pitfalls, the most serious of which are rate heterogeneity across taxa and its susceptibility to long-branch attraction (Felsenstein 1988) that can mislead phylogenetic interpretation (Eernisse 1997, Abouheif et al. 1998). As the field matured, other markers, such as mitochondrial genomes, Hox genes, the large nuclear ribosomal subunit (28S rDNA) gene, the elongation factor 1α gene, and the myosin II gene, began to be used. Additionally, the number of taxa employed in phylogenetic analyses of Metazoa increased significantly from early studies with less than 10 taxa (e.g. Halanych 1995, Bromham and Degnan 1999) to studies with over 100 taxa (e.g. Giribet et al. 2000, Eernisse and Peterson 2001). Studies that employ a target-gene approach are dependent on polymerase chain reaction (PCR) to amplify specific DNA markers for sequencing. Until recently, the use of single-copy nuclear protein-coding genes for metazoan phylogeny has been limited because of variation in intron boundaries, challenges involved in working with RNA, and the need for preexisting sequence data for primer design. As more genomic resources have become available for a number of taxa, and molecular methods have developed, phylogenetic investigations targeting multiple nuclear protein-coding genes have become more feasible (e.g. Peterson et al. 2004, Rokas et al. 2005, Bourlat et al. 2008, Paps et al. 2009). Also, highthroughput genomic approaches allowing for molecular phylogenetic data to be obtained in a PCR-independent manner have become more accessible and widespread. Studies using high-throughput sequencing have produced large amounts of data for phylogenetic analyses on growing numbers of taxa. Such approaches are typically referred to as phylogenomic approaches (Delsuc et al. 2005, Telford 2008). Most recent phylogenomic studies have made use of expressed sequence tag (EST) data. By way of a brief description, EST data are generated by extracting mRNA, reverse transcribing it to cDNA, and then sequencing a randomly selected subset of the cDNA, producing partial sequences or ‘tags.’ Data collection for most phylogenomic studies has been conducted using capillary sequencers, and by necessity has included bacterial cloning of the cDNA. As of the time of writing this manuscript, most published EST studies used capillary sequencers and relied on data produced from 1000–5000 clones. However, the development of pryosequencing technologies (e.g. 454, Illumina) has made it possible to collect very large amounts of transcriptome data at a low cost per base pair relative to capillary sequencing. The phylogenomic approach is a powerful one for studies of deep animal relationships because many of the sequences obtained belong to ‘housekeeping’ genes which are vital to the function of any given animal cell. Because such genes are usually constitutively expressed, they are likely to be recovered in a typical EST survey regardless of the source of the material. Furthermore, because

© 2011 by Taylor and Francis Group, LLC

18

Key Transitions in Animal Evolution

of their functional importance, housekeeping genes tend to be highly evolutionarily conserved, furnishing phylogenetic signal for the study of deep relationships. The first major paper using the phylogenomic approach for deep animal phylogeny was Philippe et al. (2005) in which 146 genes from 35 species were analyzed. This manuscript was followed by a series of manuscripts (e.g. Bourlat et al. 2006, Delsuc et al. 2006, Matus et al. 2006, Webster et al. 2006, Philippe et al. 2007) that added new taxa but employed nearly the same set of genes. However, other studies (e.g. Marlétaz et al. 2006, Dunn et al. 2008) have independently examined primary sequence data and have generated alternative gene sets. Before we detail some of the findings from such studies (below), we wish to outline some issues of the phylogenomic approach as currently employed. The appeal of phylogenomics stems from the fact that large amounts of genomic data corresponding to multiple genes can be brought to bear on phylogenetic issues. Additionally, the approach allows access to genetic data that would be otherwise difficult to obtain using a PCRbased approach as discussed above. However, phylogeneticists are still learning how to efficiently analyze such large data sets. Insights from previous theoretical discussions during the past 20 years focusing on, for example, effects of missing data, taxon sampling issues, signal in the data, and when to combine data have not been thoroughly addressed in a phylogenomic context. Most EST-based data sets have large amounts of missing data for any given taxon. For example, there was ~25% missing data in the matrix of Philippe et al. (2004) and ~50% missing data in the matrix of Dunn et al. (2008). While some have advocated this is not a serious problem (e.g. Philippe et al. 2004, Wiens 2006), the issue has not been adequately explored for large multigene data sets (but see Hartmann and Vision 2008). Also, some studies have had poor taxon sampling and therefore recovered artifactual results. For example, the Cephalochordata + Echinodermata clade recovered by Delsuc et al. (2006) has subsequently been shown to be an artifact of insufficient taxon sampling (Bourlat et al. 2006, further discussed below). The issue of conflicting signal in different gene partitions within a data set has received little attention in metazoan phylogenomics (but see Gatesy et al. 2007 and Wiens et al. 2008). Such analysis is commonplace for analyses of data generated using a PCRbased approach (e.g. Danforth et al. 2005, Paps et al. 2009). Lastly, one of the most represented groups of genes examined in EST-based studies is ribosomal protein genes. These genes can comprise the majority of the usable data produced by surveys of a few thousand clones but their utility has recently been questioned (e.g. Yeang and Haussler 2007, Bleidorn et al. 2009a). Because of these issues, we encourage careful consideration of the data matrix employed in phylogenomic analyses. As revealed by

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

19

discussions below, phylogenomic analyses have substantial utility and we look forward to further insights provided by such studies. Overview of Animal Phylogeny Despite holes in our understanding of animal evolutionary relationships, several hypotheses regarding higher-level metazoan relationships are now widely accepted. At the base of the animal tree are Porifera, Placozoa, Ctenophora, and Cnidaria. The relationships among these basal metazoans remain very controversial; particularly, which group has the distinction of being the basal-most animal. The existence of the clade Bilateria has been strongly supported by virtually all studies of morphology and molecules. Within Bilateria, most animals fall into three major lineages: Deuterostomia (e.g. echinoderms and chordates), Lophotrochozoa (e.g. molluscs and annelids), and Ecdysozoa (e.g. arthropods and nematodes). Exceptions include Acoela and Nemertodermatida which appear to be basal to the rest of the Bilateria and Chaetognatha which may be basal to Protostomia (Lophotrochozoa + Ecdysozoa; Marlétaz et al. 2006, Marlétaz and Le Parco 2008, but see Helmkampf et al. 2008a, Matus et al. 2006, 2007). Also, the placement of several so-called minor phyla including Orthonectida, Dicyemida, and Myxozoa on the animal tree of life is currently unclear. In the following sections, we have attempted to concisely synthesize the most recent work on animal phylogenetics and illustrate gaps in our current knowledge. Basal metazoans Relationships among the basal metazoans have received a great deal of attention recently, but despite a growing number of datasets with multiple sources of information, a consensus has not been reached. The branching order of Placozoa, Porifera (as either a monophyletic group or a paraphyletic assemblage), Ctenophora, Cnidaria, and Bilateria has been the subject of intense debate (see Cartwright and Collins 2007, Martindale 2005, Schierwater et al. 2009a). Placozoa, a phylum which has multiple lineages despite only one formally described species, Trichoplax adherens (Voigt et al. 2004, Pearse and Voigt 2007, Signorovitch et al. 2007), has been suggested to occupy every imaginable position at the base of the metazoan tree (reviewed in Schierwater et al. 2009a,b). Although placozoan mitochondrial genomes (Dellaporta et al. 2006, Signorovitch et al. 2007) and the nuclear genome of Trichoplax adherens have been sequenced (Srivastava et al. 2008), the phylogenetic affinities of Placozoa remain unresolved. Using 104 nuclear protein-coding genes from whole genomes, Srivastava et al. (2008) found

© 2011 by Taylor and Francis Group, LLC

20

Key Transitions in Animal Evolution

Trichoplax as sister to a clade of Cnidaria + Bilateria. Because this analysis was restricted to species with complete genomes, only nine taxa were sampled including one sponge, two cnidarians, and no ctenophores. In a more recent phylogenomic study with enhanced taxon sampling of basal metazoans, placozoans were found to be sister to a clade comprised of Bilateria and monophyletic Coelenterata (Cnidaria + Ctenophora) based on phylogenomic analyses (Philippe et al. 2009). Schierwater et al. (2009b) concatenated a dataset of morphological characters, mitochondrial and nuclear ribosomal gene sequences, molecular morphology, and nuclear protein-coding genes. Their analyses recovered a monophyletic Diploblasta comprised of Placozoa sister to a clade of Porifera + Coelenterata. Early molecular studies on sponges using LSU and SSU indicated that Porifera is a paraphyletic grade, with Calcerea more closely related to Eumetazoa than Demospongia (e.g. Collins 1998, Borchiellini et al. 2001, Medina et al. 2001, Cavalier-Smith and Chao 2003). This finding has been upheld using a small set of nuclear protein-coding genes (Peterson and Butterfield 2005, Sperling et al. 2007). Phylogenomic studies, however, suggest Porifera is monophyletic (Dunn et al. 2008, Philippe et al. 2009). Insights gained from poriferan mitogenomics are discussed elsewhere in this volume by Lavrov. Monophyletic or not, the phylogenetic position of Porifera has fluctuated in recent years. Investigations regarding sponge relationships were recently reviewed by Erpenbeck and Wörheide (2007). Historically, Cnidaria and Ctenophora were allied as Coelenterata but this relationship was refuted by a number of early molecular studies (see Halanych 2004). Unlike sponges and placozoans, cnidarians and ctenophores have nerve cells, true epithelial tissues, and germ layers which have been interpreted as shared derived features with Bilateria by some. Surprisingly, Dunn et al. (2008) found Ctenophora to be the basalmost metazoan phylum while Cnidaria and Porifera formed a clade basal to Bilateria. Placozoa was not included in the analysis. A more recent phylogenomic analysis by Philippe et al. (2009), however, had greatly expanded taxon sampling at the base of the metazoan tree and recovered a more traditional branching order—Porifera as the basal-most clade in Metazoa, followed by Placozoa, followed by a monophyletic Coelenterata (Ctenophora + Cnidaria). Schierwater et al.’s (2009b) combined dataset including morphology, ribosomal secondary structure, and sequence data found Placozoa, Porifera, Cnidaria, and Ctenophora to form a clade, Diploblasta, as the sister to Bilateria. Nuclear protein-coding genes suggest the myxozoan Buddenbrockia plumatellae is sister to Medusozoa within Cnidaria (Jiménez-Guri et al. 2007). Cnidarian systematics was recently reviewed by Daly et al. (2007).

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

21

Acoela and Nemertodermatida In recent years, evidence that the acoelomorph flatworms (Acoela and Nemertodermatida) are not members of Platyhelminthes, but rather are basal bilaterians, has continued to accumulate (e.g. Jondelius et al. 2002, Cook et al. 2004, Wallberg et al. 2007, Baguñà et al. 2008, Wheeler et al. 2009,). There is also a growing body of evidence suggesting that Acoelomorpha is not monophyletic; SSU/LSU + nuclear protein coding gene phylogenies have shown Acoela and Nemertodermatida as distinct lineages at the base of Bilateria with Acoela basal (Ruiz-Trillo et al. 2002, Wallberg et al. 2007, Paps et al. 2009). To date, acoels have been unstable in phylogenomic studies (Philippe et al. 2007, Dunn et al. 2008). Acoela clustered with deuterostomes in the phylogenomic analyses by Philippe et al. (2007) which was interpreted as an artifact of long branch attraction (LBA). Notably, a relationship between Acoela and Xenoturbella has been proposed based on similarities in the ultrastructure of epidermal locomotory ciliary rootlets (Lundin 1998). EST data from nemertodermatids and more slowly-evolving acoels will hopefully aid resolution. Deuterostomia Deuterostome relationships are among the best known of all animal groups (Halanych 2004, Swalla and Smith 2008). However, recent studies utilizing genomic data, particularly multigene analyses utilizing ESTs, have called into question the evolutionary history of this well-studied clade, particularly chordates. Until recently, the most commonly accepted phylogenetic hypothesis concerning chordates consisted of Tunicata as sister to Euchordata (Cephalochordata + Craniata). Phylogenomic studies have renewed interest in an alternative hypothesis in which cephalochordates are sister to a clade termed Olfactores, consisting of Tunicata and Craniata. In the EST phylogenetic study of Philippe et al. (2005), tunicates were recovered as sister to craniates, to the exclusion of cephalochordates. This study included one representative each of tunicates, cephalochordates, and echinoderms, and two vertebrate taxa. Delsuc et al. (2006) included four additional tunicate species and seven additional vertebrate species, and found tunicates clustered with craniates. Unexpectedly, the cephalochordate Branchiostoma clustered with Strongylocentrotus purpuratus, the sole echinoderm included in the analyses. This result was later shown to be due to poor taxon sampling (Bourlat et al. 2006). Olfactores, on the other hand, has been recovered in a number of phylogenomic studies utilizing different EST screening protocols as well as models of evolution (Delsuc et al. 2008, Dunn et al. 2008, Lartillot and Philippe 2008, Philippe et al. 2009). Independent support for Olfactores arrangement has come in the form of morphology (Ruppert 2005), combined rRNA and morphological data (Zrzavý et al. 1998), cadherin gene structure (Oda et al. 2004), chordate

© 2011 by Taylor and Francis Group, LLC

22

Key Transitions in Animal Evolution

fibrillar collagen genes (Wada et al. 2006), and microRNA distribution (Heimberg et al. 2008). On the other hand, analyses based on mitochondrial data (Yokobori et al. 2005, Bourlat et al. 2006) and rRNA (Mallat and Winchell 2007) recover Euchordata (Craniata + Cephalochordata) with varying degrees of support. The potential sister relationship between craniates and tunicates suggests that similarities between chordates and cephalochordates may be chordate symplesiomorphies. Tunicates are unusual in their highly derived morphologies and modified genomes and have been difficult to place phylogenetically due to long branches (Winchell et al. 2002, Zeng and Swalla, 2005, Mallatt and Winchell 2007). Whether or not the Tunicata + Craniata relationship continues to be upheld with increased taxon sampling remains to be seen. Molecular phylogenetic studies have continued to provide robust support for Ambulacraria, a clade comprised of Hemichordata and Echinodermata, first proposed by Metchnikoff in 1881 based on similarities in organization of the larval coeloms in these two groups, and formalized as a node-based name based on SSU data (Halanych 1995). Recent support for this clade has inlcuded phylogenomics (Bourlat et al. 2006, Dunn et al. 2008, Lartillot and Philippe 2008, Philippe et al. 2009), morphology (Cameron 2005, Ruppert 2005), microRNAs (Wheeler et al. 2009), and a combined analysis of ribosomal, mitochondrial and nuclear proteincoding genes (Bourlat et al. 2008). The Ambulacraria hypothesis suggests that shared morphological features between hemichordates and chordates, such as pharyngeal gill slits and the post-anal tail, were likely to have been present in the deuterostome ancestor. Recent efforts have been made to resolve relationships within hemichorates and echinoderms (Cannon and Rychel et al. 2009, Mallatt and Winchell 2007, respectively). The newly erected deuterostome phylum Xenoturbellida comprises Xenoturbella, a genus of small (< 4cm), morphologically non-descript animals (Bourlat et al. 2006). Although most analyses have found Xenoturbella sister to Ambulacraria (Bourlat et al. 2003, Bourlat et al. 2006, Bourlat et al. 2008, Dunn et al. 2008, Lartillot and Philippe 2008), phylogenetic analyses based on Hox genes (Fritzsch et al. 2008), and mitochondrial genomes (Perseke et al. 2007) have placed Xenoturbellida at the base of the deuterostomes. However, Philippe et al.’s (2007) phylogenomic study found the acoel Convoluta pulchra grouping with deuterostomes in 90% of bootstrap replicates, and despite the extremely high rate of evolution in the acoel, the authors suggest the possibility of a Xenoturbellida-Acoel relationship. Lophotrochozoa Lophotrochozoa was defined by Halanych et al. (1995) on the basis of an analysis of 18S rDNA as the last common ancestor of the three traditional lophophorate taxa (Brachiopoda, Phoronida, and Bryozoa [=Ectoprocta]),

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

23

molluscs, annelids, and all of the descendants of that common ancestor. Monophyly of Lophotrochozoa has since been supported by numerous molecular phylogenetic investigations (e.g. Anderson et al. 2004, Philippe et al. 2005, Hausdorf et al. 2007, Dunn et al. 2008, Helmkampf et al. 2008a,b). However, the interrelationships among the phyla that constitute this taxon are largely unclear. Broadly, most lophotrochozoans appear to fall into one of two major clades: Trochozoa and Platyzoa. Trochozoa (sensu Peterson and Eernisse 2001) includes Mollusca, Annelida, Nemertea, Brachiopoda, Phoronida and Entoprocta; phyla in which the presence of a trochophore larval stage is thought to be a plesiomorphy (although it has been lost or modified in some). Recent studies have greatly improved our understanding of Trochozoan phylogeny. In their phylogenomic analyses, Dunn et al. (2008) recovered a monophyletic Eutrochozoa in which Mollusca was sister to a clade comprised of Annelida, Brachiopoda, Phoronida, and Nemertea. Annelida was sister to a clade comprised of a Brachiopoda/Phoronida clade, and Nemertea. A similar topology for Eutrochozoa was recovered in a ML analysis of ribosomal proteins derived from EST data by Helmkampf et al. (2008b). Paps et al. (2009) analyzed 13 nuclear protein-coding genes and also recovered a monophyletic Trochozoa (albeit with weak support). Molluscs are an ancient group of morphologically diverse trochozoans whose class-level relationships have proven difficult to reconstruct. To date, investigations of molluscan relationships have been unable to resolve most deep branches. The most significant results of these studies include the rejection of the Diasoma hypothesis (which allies scaphopods and bivalves; Passamaneck et al. 2004) and evidence for a clade of cephalopods and scaphopods (Steiner and Dreyer 2002, 2003). The phylogenetic position of Monoplacophora was recently addressed by Wilson et al. (2009). Molluscan phylogenetics were recently reviewed in Phylogeny and Evolution of the Mollusca (Ponder and Lindberg, eds. 2008). Recent molecular phylogenetic investigations have radically altered our understanding of Annelida by demonstrating that Clitellata is within ‘Polychaeta’ and several taxa formerly thought to be distinct phyla (Pogonophora [now Siboglinidae], Echiura, and Sipuncula) and are actually within the annelid radiation (McHugh 1997, Colgan et al. 2006, Passamaneck and Halanych 2006, Rousset et al. 2007, Struck et al. 2007). The annelid affinities of Myzostomida, a group of unusual marine parasites, are still being debated. Recent studies of myzostomid anatomy (Eeckhaut and Lanterbecq 2005, Lanterbecq et al. 2008), mitochondrial genome information, nuclear rDNA + protein-coding genes (Bleidorn et al. 2007, 2009a), and Hox genes (Bleidorn et al. 2009b) support a close relationship between Myzostomida and Annelida. See Zrzavý et al. (2009) for the most recent phylogenetic investigation of Annelida.

© 2011 by Taylor and Francis Group, LLC

24

Key Transitions in Animal Evolution

Brachiopods, phoronids, and bryozoans (= ectoprocts) have traditionally been allied as the Lophophorata (Nielsen 1985, 2001) and while they all possess a horseshoe-shaped ring of hollow tentacles termed a lophophore, there are structural differences (Halanych 1996). However, recent work has mostly supported the monophyly of Brachiopoda and Phoronida to the exclusion of Bryozoa (e.g. Passamaneck and Halanych 2006, Anderson et al. 2004, Dunn et al. 2008, Helmkampf et al. 2008b). The phylogenetic affinities of Bryozoa are presently unclear. Phylogenomic analyses by Hausdorf et al. (2007) and Helmkampf et al. (2008b) recovered a well-supported sister relationship between the ectoproct Flustra and the entoproct (= kamptozoan) Barentsia but this is surprising given morphology (Wanninger 2009). Other studies place Entoprocta as the sister taxon of Cycliophora (Passamaneck and Halanych 2006). See Fuchs et al. (2009) for the most up-to-date phylogeny of Bryozoa. Platyzoa comprises most of the remainder of Lophotrochozoa and is usually defined to include Platyhelminthes (excluding Acoelomorpha), Gastrotricha, Rotifera (including Acanthocephala), Gnathostomulida, and Micrognathozoa (Garey et al. 1996, Garey and Schmidt-Rhaesa 1998, Giribet et al. 2000, Passamaneck and Halanych et al. 2006). Virtually nothing is known about the relationships among the taxa that comprise Platyzoa and it was even suggested by Dunn et al. (2008) that Platyzoa is an artifact of long-branch attraction. Ecdysozoa Ecdysozoa comprises eight phyla which are united by the molting of their cuticle. Aguinaldo et al. (1997) using SSU rDNA provided the first molecular support for this grouping. Since then, several lines of evidence have supported the existence of this clade (reviewed by Halanych 2004, Telford et al. 2008). Within Ecdysozoa, three major clades are typically recognized: Scalidophora (Priapulida, Kinorhyncha, and Loricifera), Nematoida (Nematoda and Nematomorpha), and Panarthropoda (Arthropoda, Tardigrada, and Onychophora). The evolution of this clade has recently been reviewed by Telford et al. (2008), Budd and Telford (2009) and Edgecombe (2009). The clade Nematoida (Nematoda + Nematomorpha) has been supported by SSU rDNA and morphology (Peterson and Eernisse 2001), SSU rDNA and LSU rDNA (e.g. Petrov and Vladychenskaya 2005, Mallatt and Giribet 2006), concatenated nuclear, ribosomal and mitochondrial genes (Bourlat et al. 2008), and phylogenomics (Dunn et al. 2008). Some studies have found other taxa pairing with nematoids, including Loricifera and Tardigrada, with varying degrees of support (see further discussion below), but the nematomorph-nematode relationship seems to be robust. Most molecular studies have supported a Priapulid + Kinorhynch clade (Petrov and Vladychenskaya 2005, Mallat and Giribet 2006, Dunn et

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

25

al. 2008), but the inclusion of Loricifera in Scalidophora has been called into question by a molecular phylogeny by Sørensen et al. (2008), whose results suggested a possible relationship between Loricifera and Nematomorpha. Embryological studies have been inconclusive as to the relationships within Scalidophora—the developmental patterns of Priapulids (Wennberg et al. 2009), Kinorhynchs (Kozloff 2007), and Loriciferans (Gad 2005) do not appear to closely resemble each other. Scalidophora and Nematoida have been united in a clade called Cyloneuralia or Introverta, with the names referring to their circum-oral brain and eversible anterior introvert, respectively. Molecular analyses based on SSU rDNA and LSU rDNA have not recovered this clade (Mallat and Giribet 2006, Sørensen et al. 2008), but Dunn et al. (2008) did recover Cycloneuralia (Loricifera was not included in the analysis). Panarthropoda was hypothesized on the basis of morphological characters shared among Arthropoda, Onychophora, and Tardigrada. Evolution of this group, including information on several recently discovered stem-group fossil taxa, was recently reviewed by Budd and Telford (2009) and Edgecombe (2009). The immediate sister taxon of Arthropoda has been a long-standing question, but two recent phylogenomic studies have supported a sister taxon relationship between Arthropoda and Onychophora (Roeding et al. 2007, Dunn et al. 2008). The phylogenetic position of Tardigrada, however, remains unclear. Roeding et al. (2007) found tardigrades to be more closely related to Nematoda than Arthropoda or Onychophora. However, in the 64-taxon analysis by Dunn et al., tardigrades were recovered as sister to Arthropoda + Onychophora, consistent with the Panarthropoda hypothesis. Although numerous morphological features suggest a close relationship between tardigrades and arthropods, several cycloneuralian-like features of tardigrades may be in need of re-evaluation (reviewed by Edgecombe 2009). Arthropoda has traditionally been divided into four major lineages: Chelicerata, Myriapoda, Crustacea, and Hexapoda. Classical hypotheses regarding arthropod phylogeny are reviewed by Budd and Telford (2009). More recently, evidence suggesting that Hexapoda is nested within Crustacea has come from molecular phylogenetic studies (Richter 2002, Regier et al. 2005, Regier et al. 2008) as well as morphological studies (e.g. Ungerer and Scholtz 2008). This clade of crustaceans and hexapods has been termed Pancrustacea or Tetraconata. The phylogenetic relationships among Pancrustacea, Myriapoda and Chelicerata remain uncertain. There is evidence for a clade of Pancrustacea + Myriapoda (=Mandibulata; Edgecombe et al. 2003, Scholtz and Edgecombe 2006, Mallatt and Giribet 2006) or, alternatively, a clade of Chelicerata + Myriapoda (=Paradoxopoda or Myriochelata; Negrisolo et al. 2004, Pisani et al. 2004). Support for both hypotheses has varied; recent studies including Dunn et al. (2008) found

© 2011 by Taylor and Francis Group, LLC

26

Key Transitions in Animal Evolution

support for Paradoxopoda while the others such as Bourlat et al. (2008) and Regier et al. (2008) were more consistent with Mandibulata. Conclusions Despite many recent advances in the field of animal phylogenetics, a number of daunting questions remain unanswered, including the relationships among the basal metazoans, the branching order of the three major lineages of Arthropoda, relationships within Lophotrochozoa, and the placement of many so-called minor groups such as Chaetognatha. The phylogenetic tree in Fig. 1 presents a conservative view of our current understanding of animal evolutionary relationships. Polytomies

Fig. 1. Current understanding of animal phylogeny. This tree is conservatively drawn to illustrate major concepts inferred from recent studies as described in the text. The tree is color-coded: ‘basal metazoans’ are purple, deuterostomes are red, ecdysozoans are blue, and lohpotrochozoans are green. *The monophyly of Porifera has been questioned, but for the sake of simplicity it is drawn as a single lineage here (see text). Color image of this figure appears in the color plate section at the end of the book.

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

27

(unresolved nodes) within Arthropoda, Lophotrochozoa, and at the base of the tree represent forthcoming challenges to phylogeneticists. Recent developments in the fields of phylogenomics, gene expression analysis, pyrosequencing, microscopy and 3D visualization, and paleontology will undoubtedly continue to help refine our understanding of the animal tree of life, thereby broadly informing studies of metazoan evolution. References Abouheif, E. and R. Zardoya, and A. Meyer. 1998. Limitations of metazoan 18S rRNA sequence data: implications for reconstructing a phylogeny of the animal kingdom and inferring the reality of the Cambrian explosion. J. Mol. Evol. 47: 394–405. Aguinaldo, A.M.A. and J.M. Turbeville, L.S. Linford, M.C. Rivera, J.R. Garey, R. Raff, and J.A. Lake. 1997. Evidence for a clade of nematodes, arthropods and other moulting animals. Nature 387: 489–493. Anderson, F.E. and A.J. Cordoba, and M. Thollesson. 2004. Bilaterian phylogeny based on analyses of a region of the sodium-potassium ATPase alpha subunit gene. Mol. Evol. 58: 252–268. Baguñà, J. and P. Martinez, J. Paps, and M. Riutort. 2008. Back in time: a new systematic proposal for the Bilateria. Phil. Trans. R. Soc. B 2008 363: 1481–1491. Bleidorn, C. and I. Eeckhaut, L. Podsiadlowski, N. Schult, D. McHugh, K.M. Halanych, M.C. Milinkovitch, and R. Tiedemann. 2007. Mitochondrial genome and nuclear sequence data support Myzostomida as part of the annelid radiation. Mol. Biol. Evol. 24: 1690– 1701. Bleidorn, C. and L. Podsiadlowski, M. Zhong, I. Eeckhaut, S. Hartmann, K.M. Halanych, and R. Tiedemann. 2009a. On the phylogenetic position of Myzostomida: can 77 genes get it wrong? BMC Evol. Biol. 9: 150. Bleidorn, C. and D. Lanterbecq, I. Eeckhaut, and R. Tiedemann. 2009b. A PCR survey of Hox genes in the myzostomid Myzostoma cirriferum. Dev. Genes Evol. 219: 211–216. Borchiellini, C. and M. Manuel, E. Alivon, N. Boury-Esnault, J. Vacelet, and Y. Le Parco. 2001. Sponge paraphyly and the origins of metazoa. J. Evol. Biol. 14: 171–179. Bourlat, S.J. and C. Nielsen, A.E. Lockyer, D.T.J. Littlewood and M.J. Telford, 2003. Xenoturbella is a deuterostome that eats molluscs. Nature 424: 925–928. Bourlat, S.J. and T. Juliusdottir, C.J. Lowe, R. Freeman, J. Aronowicz, M. Kirschner, E.S. Lander, M. Thorndyke, H. Nakano, and A.B. Kohn. 2006. Deuterostome phylogeny reveals monophyletic chordates and the new phylum Xenoturbellida. Nature 444: 85–88. Bourlat, S.J. and C. Nielsen, A.D. Economou, and M.J. Telford. 2008. Testing the new animal phylogeny: A phylum level molecular analysis of the animal kingdom. Mol. Phylogenet. Evol. 49: 23–31. Bromham, L.D. and B.M. Degnan. 1999. Hemichordates and deuterostome evolution: robust molecular phylogenetic support for a hemichordate + echinoderm clade. Evol. Dev. 1: 168–171. Budd, B. and M.J. Telford. 2009. The origin and evolution of arthropods. Nature 452: 812–817. Cameron, C.B. 2005. A phylogeny of the hemichordates based on morphological characters. Can. J. Zool. 83: 196–215. Cannon, J.T. and A.L. Rychel, H. Eccleston, K.M. Halanych, and B.J. Swalla. 2009. Molecular phylogeny of Hemichordata, with updated status of deep-sea enteropneusts. Mol. Phylogenet. Evol. 52: 17–24. Cartwright, P. and A. Collins. 2007. Fossils and phylogenies: Integrating multiple lines of evidence to investigate the origin of early major metazoan lineages. Int. Comp. Biol. 47: 744–751.

© 2011 by Taylor and Francis Group, LLC

28

Key Transitions in Animal Evolution

Cavalier-Smith T. and E.E. Chao. 2003. Phylogeny of Choanozoa, Apusozoa, and other Protozoa and early eukaryote megaevolution. J. Mol. Evol. 56: 540–563. Colgan, D.J. and P.A. Hutchings and M. Braune. 2006. A multigene framework for polychaete phylogenetic studies. Org. Div. and Evol. 2: 220–235. Collins, A.G. 1998. Evaluating multiple alternative hypotheses for the origin of Bilateria: An analysis of 18S rRNA molecular evidence. Proc. Natl. Acad. Sci. USA 95: 15458–15463. Cook, C.E. and E. Jiménez, M. Akam, and E. Saló. 2004. The Hox gene compliment of acoel flatworms, a basal bilaterian clade. Evol. Dev. 6: 154–163. Daly, M. and M.R. Brugler, P. Cartwright, A.G. Collins, M.N. Dawson, D.G. Fautin, S.C. France, C.S. McFadden, D.M. Opresko, E. Rodriguez, S.L. Romano, and J.L. Stake. 2007. The phylum Cnidaria: A review of phylogenetic patterns and diversity 300 years after Linnaeus. Zootaxa 1668: 127–182. Danforth, B.N. and C.P. Lin, and J. Fang. 2005. How do insect nuclear ribosomal genes compare to protein-coding genes in phylogenetic utility and nucleotide substitution patterns? Syst. Entomol. 30: 549–562. Dellaporta, S.L. and A. Xu, S. Sagasser, W. Jakob, M.A. Moreno, L.W. Buss, and B. Schierwater. 2006. Mitochondrial genome of Trichoplax adhaerens supports Placozoa as the basal lower metazoan phylum. Proc. Natl. Acad. Sci. USA 103: 8751–8756. Delsuc, F. and H. Brinkmann, and H. Philippe. 2005. Phylogenomics and the reconstruction of the tree of life. Nature Rev. Genet. 6: 361–375. Delsuc, F. and H. Brinkmann, D. Chourrout, and H. Philippe. 2006. Tunicates and not cephalochordates are the closest living relatives of vertebrates. Nature 439: 965–968. Delsuc, F. and G. Tsagkorgeorga, N. Lartillot, and H. Philippe. 2008. Additional molecular support for the new chordate phylogeny. Genesis 46: 592–604. Dunn, C.W. and A. Hejnol, D.Q. Matus, K. Pang, W.E. Browne, S.A. Smith, E. Seaver, G.W. Rouse, M. Obst, G.D. Edgecombe, M.V. Sørensen, S.H.D. Haddock, A. Schmidt-Rhaesa, A. Okusu, R.M. Kristensen, W.C. Wheeler, M.Q. Martindale, and G. Giribet. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–749. Edgecombe, G.D. and S. Richter, and G.D.F. Wilson. 2003. The mandibular gnathal edges: homologous structures throughout Mandibulata. J. Afr. Invertebr. 44: 115–135. Edgecombe, G.D. 2009. Palaeontological and molecular evidence linking arthropods, onychophorans, and other Ecdysozoa. Evol. Edu. Outreach 2: 178–190. Eeckhaut, I. and D. Lanterbecq. 2005. Myzostomida: A review of the phylogeny and ultrastructure. Hydrobiologia 535: 253–275. Eernisse, D.J. Arthropod and annelid relationships re-examined. pp. 43–56. In: R.A. Fortey and R.H. Thomas. [eds.] 1997. Arthropod Relationships. Chapman and Hall, London. Eernisse, D.J. and K.J. Peterson. 2001. Animal phylogeny and the ancestry of bilaterians: inferences from morphology and 18s rDNA gene sequences. Evol. Dev. 3: 170–205. Eernisse, D.J. and K.J. Peterson. 2004. The history of animals. pp. 197–208. In: J. Cracraft and M.J. Donoghue. [eds.] 2004. Assembling the Tree of Life. Oxford Univ. Press, New York. Erpenbeck, D. and G. Wörheide. 2007. On the molecular phylogeny of sponges. Zootaxa 1668: 107–126. Felsenstein, J. 1988. Phylogenies from molecular sequences: inference and reliability. Annu. Rev. Genet. 22: 521–565. Field, K.G. and G.J. Olsen, D.J. Lane, S.J. Giovannoni, and M.T. Ghiselin, et al. 1988. Molecular phylogeny of the animal kingdom. Science 239: 748–753. Fritzsch, G. and M.U. Böhme, M. Thorndyke, H. Nakano, O. Israelsson, T. Stach, M. Schlegel, T. Hankeln, and P.F. Stadler. 2008. PCR Survey of Xenoturbella bocki Hox genes. J. Exp. Zool. Mol. Dev. Evol. 310B: 278–284. Fuchs, J. and M. Obst, and P. Sundberg. 2009. The first comprehensive molecular phylogeny of Bryozoa (Ectoprocta) based on combined analyses of nuclear and mitochondrial genes. Mol. Phylogenet. Evol. 52(1): 225–233..

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

29

Gad, G. 2005. A parthenogenetic, simplified adult in the life cycle of Pliciloricus pedicularis sp. n. (Loricifera) from the deep sea of the Angola Basin (Atlantic). Organisms Diversity and Evol. 5: 77–103. Garey, J.R. and T.J. Near, M.R. Nonnemacher, and S.A. Nadler. 1996. Molecular evidence for Acanthocephala as a subtaxon of Rotifera. J. Mol. Evol. 43: 287–292. Garey, J.R. and A. Schmidt-Rhaesa. 1998. The essential role of “minor” phyla in molecular studies of animal evolution. Am. Zool. 38: 907–917. Gatesy, J. and R. DeSalle, and N. Wahlberg. 2007. How many genes should a systematist sample? Conflicting insights from a phylogenomic matrix characterized by replicated incongruence. Syst. Biol. 56: 355–363. Giribet, G. and D.L. Distel, M. Polz, W. Sterrer, and W.C. Wheeler. 2000. Triploblastic relationships with emphasis on the acoelomates and the position of Gnathostomulida, Cycliophora, Plathelminthes, and Chaetognatha: A combined approach of 18S rDNA sequences and morphology. Syst. Biol. 49: 539–562. Giribet, G. and C.W. Dunn, G.D. Edgecombe, and G.W. Rouse. 2007. A modern look at the animal tree of life. Zootaxa 1668: 61–79. Halanych, K.M. 1995. The phylogenetic position of pterobranch hemichordates based on 18s rDNA sequence data. Mol. Phylogenet. and Evol. 4: 72–76. Halanych, K.M. and J.D. Bacheller, A.M.A. Aguinaldo, S.M. Liva, D.M. Hillis, and J.A. Lake. 1995. Evidence from 18S ribosomal DNA that the lophophorates are protostome animals. Science 267: 1641–1643. Halanych, K.M. 1996. Convergence in the feeding apparatuses of lophophorates and pterobranch hemichordates revealed by 18S rDNA. Biol. Bull. 190: 1–5. Halanych, K.M. 1999. Metazoan phylogeny and the shifting comparative framework. pp. 3–7. In: Roubos, Wendelaar, Bonga, Vaudry, and De Loof. [eds.] Recent Developments in Comparative Endocrinology and Neurobiology. Shaker Publishing. Maastricht, The Netherlands. Halanych and K.M. 2004. The new view of animal phylogeny. Annu. Rev. Ecol. Evol. Syst. 35: 229–256. Hartmann, S. and T.J. Vision. 2008. Using ESTs for phylogenomics: Can one accurately infer a phylogenetic tree from a gappy alignment? BMC Evol. Biol. 8: 95. Hausdorf, B. and M. Helmkampf, A. Meyer, A. Witek, H. Herlyn, I. Bruchhaus, T. Hankeln, T.H. Struck, and B. Lieb. 2007. Spiralian phylogenomics supports the resurrection of Bryozoa comprising Ectoprocta and Entoprocta. Mol. Biol. and Evol. 24: 2723. Heimberg, A.M. and L.F. Sempere, V.N. Moy, P.C.J. Donoghue, and K.J. Peterson. 2008. MicroRNAs and the advent of vertebrate morphological complexity. Proc. Natl. Acad. Sci. USA 105: 2946–2950. Helmkampf, M. and I. Bruchhaus, and B. Hausdorf. 2008a. Multigene analysis of lophophorate and chaetognath phylogenetic relationships. Mol. Phylogenet. Evol. 46: 206–214. Helmkampf, M. and I. Bruchhaus, and Hausforf, B. 2008b. Phylogenomic analyses of lophophorates (brachiopods, phoronids and bryozoans) confirm the Lophotrochozoa concept. Proc. R. Soc. B. 275: 1927–1933. Hyman, L.H. 1959. The Invertebrates: Smaller Coelomate Groups. McGraw-Hill, New York. Jondelius, U. and I. Ruiz-Trillo, J. Baguñà, and M. Riutort. 2002. The Nemertodermatida are basal bilaterians and not members of the Platyhelminthes. Zool. Script. 31: 201–215. Kozloff, E.N. Stages of development, from first cleavage to hatching, of an Echinoderes (Phylum Kinorhyncha: Class Cyclorhagida). Cahiers de Biologie Marine 48: 199–206. Lanterbecq, D. and C. Bleidorn, S. Michel, and I. Eeckhaut. 2008. Locomotion and find structure of parapodia in Myzostoma cirriferum (Myzostomida). Zoomorphology 127: 59–68. Lartillot, N. and H. Philipe. 2004. A Bayesian mixture model for across-site heterogeneities in the amino-acid replacement process. Mol. Biol. Evol. 21: 1095–1109.

© 2011 by Taylor and Francis Group, LLC

30

Key Transitions in Animal Evolution

Lartillot, N. and H. Brinkman, and H. Philippe. 2007. Suppression of long-branch attraction artifacts in the animal phylogeny using a site-heterogeneous model. BMC Evol. Biol. 7 (Suppl.1): S4. Lartillot, N. and H. Philippe. 2008. Improvement of molecular phylogenetic inference and the phylogeny of Bilateria. Phil. Trans. R. Soc. B. 363: 1463–1472. Lundin, K. 1998. The epidermal ciliary rootlets of Xenoturbella bocki (Xenoturbellida) revisited: new support for a possible kinship with the Acoelomorpha (Platyhelminthes). Zool. Script. 27: 263–270. Mallatt, J. and G. Giribet. 2006. Further use of nearly complete 28S and 18S rRNA genes to classify Ecdysozoa: 37 more arthropods and a kinorhynch. Mol. Phylogenet. Evol. 40: 772–794. Mallat, J. and C.J. Winchell. 2007. Ribosomal RNA genes and deuterostome phylogeny revisted: More cyclostomes, elasmobranchs, reptiles, and a brittle star. Mol. Phylogenet. Evol. 43: 1005–1022. Marlétaz, F. and E. Martin, Y. Perez, D. Papillon, X. Caubit, C.J. Lowe, B. Freeman, L. Fasano, C. Dossat, and P. Wincker. 2006. Chaetognath phylogenomics: a protostome with deuterostome-like development. Curr. Biol. 16: R577–578. Marlétaz, F. and Y. Le Parco. 2008. Careful with understudied phyla: The case of chaetognath. BMC Evol. Biol. 8: 251. Martindale, M.Q. 2005. The evolution of metazoan axial properties. Nature Rev. Genet. 6: 917–927. Matus, D.Q. and R.R. Copley, C.W. Dunn, A. Hejnol, H. Eccleston, K.M. Halanych, M.Q. Martindale, and M.J. Telford. 2006. Broad taxon and gene sampling indicate that chaetognaths are protostomes. Curr. Biol. 16: R575–576. Matus, D.Q. and K.M. Halanych, and M.Q. Martindale. 2007. The Hox gene complement of a pelagic chaetognath, Flaccisagitta enflata. Int. Comp. Biol. 47: 854–864. McHugh, D. 1997. Evidence that echiurans and pogonophorans are derived annelids. Proc. Natl. Acad. Sci. USA 94: 8006–8009. Medina, M. and A.G. Collina, J.D. Silberman, and M.L. Sogin. 2001. Evaluating hypotheses of basal animal phylogeny using complete sequences of large and small subunit rRNA. Proc. Natl. Acad. Sci. USA 98: 9707–9712. Metschnikoff, V.E. 1881. Über die systematische Stellung von Balanoglossus. Zool. Anz. 4: 139–157. Negrisolo, E. and A. Minelli, and G. Valle 2004. The mitochondrial genome of the house centipede Scutigera and the monophyly versus paraphyly of myriapods. Mol. Biol. Evol. 21: 770–780. Nielsen, C. 1985. Animal phylogeny in the light of the trochaea theory. Biol. J. Linn. Soc. 25: 243–299. Nielsen, C. 2001. Animal Evolution: Interrelationships of the Living Phyla. Oxford University Press, Oxford. Oda, H. and Y. Akiyama-Oda, and S. Zhang. 2004. Two classic cadherin-related molecules with no cadherin extracellular repeats in the cephalochordate amphioxus: distinct adhesive specificities and possible involvements in the development of multicelllayered structures. J. Cell Sci. 117: 2757–2767. Paps, J. and B. Baguñà, and M. Riutort. 2009. Bilaterian phylogeny: A broad sampling of 13 nuclear genes provides a new Lophotrochozoa phylogeny and supports a paraphyletic basal Acoelomorpha. Mol. Biol. Evol. 26: 2397–2406. Passamaneck, Y.J. and C. Schander, and K.M. Halanych. 2004. Investigation of molluscan phylogeny using large-subunit and small-subunit nuclear rRNA sequences. Mol. Phylogenet. Evol. 32: 25–38. Passamaneck, Y.J. and K.M. Halanych. 2006. Lophotrochozoan phylogeny assessed with LSU and SSU data: Evidence of lophophorate polyphyly. Mol. Phylogenet. Evol. 40: 20–28. Pearse, V.B. and O. Voigt. 2007. Field biology of placozoans (Trichoplax): distribution, diversity, biotic interactions. Int. Comp. Biol. 47: 677–692.

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

31

Perseke, M. and T. Hankeln, B. Weich, G. Fritzsch, P.F. Stadler, O. Israelsson, D. Bernhard, and M. Schlegal. 2007. The mitochondrial DNA of Xenoturbella bocki: genomic architecture and phylogenetic analysis. Theory Biosci. 126: 35–42. Peterson, K.J. and D.J. Eernisse. 2001. Animal phylogeny and the ancestry of bilaterians: inferences from morphology at 18s rDNA gene sequences. Evol. Dev. 3: 170–205. Peterson, K.J. and J.B. Lyons, K.S. Nowak, C.M. Takacs, M.J. Wargo, and M.A. McPeek. 2004. Estimating metazoan divergence times with a molecular clock. Proc. Natl. Acad. Sci. USA 101: 6536–6541. Peterson, K.J. and N.J. Butterfield. 2005. Origin of the Eumetazoa: testing ecological predictions of molecular clocks against the Proterozoic fossil record. Proc. Natl. Acad. Sci. USA 102: 9547–9552. Petrov, N.B. and N.S. Vladychenskaya. 2005. Phylogeny of molting protostomes (Ecdysozoa) as inferred from 18S and 28S rRNA gene sequences. Mol. Biol. 39: 503–513. Philippe, H. and E.A. Snell, E. Bapteste, P. Lopez, P.W.H. Holland, and D. Casane. 2004. Phylogenomics of eukaryotes: impact of missing data on large alignments. Mol. Biol. Evol. 21: 1740–1752. Philippe, H. and N. Lartillot, and H. Brinkman. 2005. Multigene analyses of bilaterian animals corroborate the monophyly of Ecdysozoa, Lophotrochozoa, and Protostomia. Mol. Biol. Evol. 22: 1246–1253. Philippe, H. and H. Brinkmann, P. Martinez, M. Riutort, and J. Baguña. 2007. Acoel flatworms are not platyhelminthes: Evidence from phylogenomics. PLoS ONE 2: e717. Philippe, H. and R. Derelle, P. Lopez, K. Pick, C. Borchiellini, N. Boury-Esnault, J. Vacelet, E. Renard, E. Houliston, E. Quéinnec, C. Da Silva, P. Wincker, H. Le Guyader, S. Leys, D.J. Jackson, F. Schreiber, D. Erpenbeck, B. Morgenstern, G. Wörheie, and M. Manuel. 2009. Phylogenomics revives traditional views on deep animal relationships. Curr. Biol. 19: 706–712. Pisani, D. and L.L. Poling, M. Lyons-Weiler, and S.B. Hedges. 2004. The colonization of land by animals: molecular phylogeny and divergence among arthropods. BMC Biol. 2: 1. Ponder, W.F. and D.R. Lindberg. 2009. Phylogeny and Evolution of the Mollusca. University of California Press. Berkeley and Los Angeles, CA. Regier, J.C. and J.W. Shultz, and R.E. Kambic. 2005. Pancrustacean phylogeny: Hexapods are terrestrial crustaceans and maxillopods are not monophyletic. Proc. R. Soc. B. 272: 395–401. Regier, J.C. and J.W. Shultz, A.R.D. Ganley, A. Hussey, D. Shi, B. Ball, A. Zwick, J.E. Stajich, M.P. Cummings, J.W. Martin, and C.W. Cunningham. 2008. Resolving arthropod phylogeny: exploring phylogenetic signal within 41 kb of protein-coding nuclear gene sequence. Syst. Biol. 57: 920–938. Richter, S. 2002. The Tetraconata concept: hexapod-crustacean relationships and the phylogeny of Crustacea. Org. Divers. Evol. 2: 217–237. Roeding, F. and S. Hagner-Holler, H. Ruhberg, I. Ebersberger, A. von Haeseler, M. Kube, R. Reinhardt, and T. Burmester. 2007. EST sequencing of Onychophora and phylogenomic analysis of Metazoa. Mol. Phylogenet. Evol. 45: 942–951. Rokas, A. and D. Krüger, and S.B. Carroll. 2005. Animal evolution and the molecular signature of radiations compressed in time. Science 310: 1933–1938. Rousset, V. and F. Pleijel, G.W. Rouse, C. Erséus, and M.E. Siddall. 2007. A molecular phylogeny of annelids. Cladistics 23: 41–63. Ruiz-Trillo, I. and M. Riutort, T.J. Littlewood, E.A. Herniou, and J. Baguña. 1999. Acoel flatworms: earliest extant bilaterian metazoans, not members of Platyhelminthes. Science 283: 1919–1923. Ruiz-Trillo, I. and J. Paps, M. Loukota, C. Ribera, U. Jondelius, J. Baguña, and M. Riutort. 2002. A phylogenetic analysis of myosin heavy chain type II sequences corroborates that Acoela and Nemertodermatida are basal bilaterians. Proc. Natl. Acad. Sci. USA 99: 11246–11251.

© 2011 by Taylor and Francis Group, LLC

32

Key Transitions in Animal Evolution

Ruppert, E.E. 2005. Key characters uniting hemichordates and chordates: homologies or homoplasies? Can. J. Zool. 83: 8–23. Schierwater, B. and D. de Jong, and R. DeSalle. 2009a. Placozoa and the evolution of metozoa and intrasomatic cell differentiation. Int. J. Biochem. Cell Biol. 41: 370–379. Schierwater, B. and M. Eitel, W. Jakob, H. Osigus, H. Hadrys, S.L. Dellaporta, S. Kolokotronis, and R. DeSalle. 2009b. Concatenated analysis sheds light on early metazoan evolution and fuels a modern “Urmetazoon” hypothesis. PLoS Biol. 7: e1000020. Signorovitch, A.Y. and L.W. Buss, and S.L. Dellaporta. 2007. Comparative genomics of large mitochondria in placozoans. PLoS Genet. 3: e13. Sørensen, M.V. and M.B. Hebsgaard, I. Heiner, H. Glenner, E. Willerslev, and R.M. Kristensen. 2008. New data from an enigmatic phylum: evidence from molecular sequence data supports a sister-group relationship between Loricifera and Nematomorpha. J. Zool. Syst. Evol. Res. 46: 231–239. Sperling, E.A. and D. Pisani, and K.J. Peterson. Poriferan paraphyly and its implications for Precambrian palaeobiology. In: P. Vickers-Rich and P. Komarower [eds.] 2007. The Rise and Fall of the Ediacaran Biota. The Geological Society of London, London. Srivastava, M. and E. Begovic, J. Chapman, N.H. Putnam, U. Hellstein, T. Kawashima, A. Kuo, T. Mitros, A. Salamov, M.L. Carpenter, A.Y. Signorovitch, M.A. Moreno, K. Kamm, J. Grimwood, J. Schmutz, H. Shapiro, I.V. Grigoriev, L.W. Buss, B. Schierwater, S.L. Dellaporta, and D.S. Rokhsar. The Trichoplax genome and the nature of placozoans. Nature 454: 955–960. Steiner, G. and H. Dreyer. 2002. Cephalopoda and Scaphopoda are sister taxa: an evolutionary scenario. Zoology 105: 95. Steiner, G. and H. Dreyer. 2003. Molecular phylogeny of Scaphopoda (Mollusca) inferred from 18S rDNA sequences: support for a Scaphopoda-Cephalopoda clade. Zool. Scr. 32: 343–356. Struck, T.H. and N. Schult, T. Kusen, E. Hickman, C. Bleidorn, D. McHugh, and K.M. Halanych. 2007. Annelid phylogeny and the status of Sipuncula and Echiura. BMC Evol. Biol. 7: 57. Swalla, B.J. and A.B. Smith. 2008. Deciphering deuterostome phylogeny: molecular, morphological and palaeontological perspectives. Phil. Trans. R. Soc. B 363: 1557–1568. Telford, M.J. 2008. Resolving animal phylogeny: A sledgehammer for a tough nut? Dev. Cell 14: 457–459. Telford, M.J. and S.J. Bourlat, A. Economou, D. Papillon, and O. Rota-Stabelli. 2008. The evolution of the Ecdysozoa. Phil. Trans. R. Soc. B 363: 1529–1537. Ungerer, P. and G. Scholtz. 2007. Filling the gap between identified neuroblasts and neurons in crustaceans adds new support for Tetraconata. Proc. R. Soc. B. 275: 369–376. Voigt, O. and A.G. Collins, V. Buchsbaum Pearse, J.S. Pearse, A. Ender, H. Hadrys, and B. Schierwater. Placozoa—no longer a phylum of one. Curr. Biol. 14: R944–R945. Wada, H. and M. Okuyama, N. Satoh, and S. Zhang. 2006. Molecular evolution of fibrillar collagen in chordates, with implications for the evolution of vertebrate skeletons and chordate phylogeny. Evol. Dev. 8: 370–377. Wallberg, A. and M. Curini-Galletti, A. Ahmadzadeh, and U. Jondelius. 2007. Dismissal of Acoelomorpha: Acoela and Nemertodermatida are separate early bilaterian clades. Zool. Script. 36: 509–523. Wanninger, A. 2009. Shaping the things to come: Ontogeny of lophotrochozoan neuromuscular systems and the Tetraneuralia concept. Biol. Bull. 216: 293–206. Webster, B.L. and R.R. Copley, R.A. Jenner, J.A. Mackenzie-Dodds, S.J. Bourlat, O. Rota-Stabelli, D.T.J. Littlewood, and M.J. Telford. 2006. Mitogenomics and phylogenomics reveal priapulid worms as extant models of the ancestral Ecdysozoan. Evol. Dev. 8: 502–510. Wennberg, S.A. and R. Janssen, and G.E. Budd. 2009. Hatching and earliest larval stages of the priapulid worm Priapulus caudatus. Invertebr. Biol. 128: 157–171. Wheeler, B.M., A.M. Heimberg, V.N. Moy, E.A. Sperling, T.W. Holstein, S. Heber, and K.J. Peterson. 2009. The deep evolution of metazoan microRNAs. Evol. Dev. 11: 50–68.

© 2011 by Taylor and Francis Group, LLC

Metazoan Phylogeny

33

Willmer, P. 1990. Invertebrate Relationships, Patterns in Animal Evolution. Cambridge Univ. Press, New York. Wiens, J.J. 2006. Missing data and the design of phylogenetic analyses. Biomed. Infomat. 39: 34–42. Wiens, J.J. and C.A. Kuczynski, S.A. Smith, D.G. Mulcahy, J.W. Sites Jr., T.M. Townsend, and T.W. Reeder. 2008. Branch lengths support and congruence testing the phylogenomic approach with 20 nuclear loci in snakes. Syst. Biol. 57: 420–431. Winchell, C.J. and J. Sullivan, C.B. Cameron, B.J. Swalla, and J. Mallatt. 2002. Evaluating hypotheses of deuterostome phylogeny and chordate evolution with new LSU and SSU ribosomal DNA data. Mol. Biol. Evol. 19: 762–776. Wilson, N.G. and G.W. Rouse, and G. Giribet. 2009. Assessing the molluscan hypothesis Serialia (Monoplacophora + Polyplacophora) using novel molecular data. Mol. Phylogenet. Evol. 54(1): 187–193.. Yeang, C. and D. Haussler. 2007. Detecting coevolution in and among protein domains. PLoS Comp. Biol. 3: e211. Yokobori, S. and T. Oshima, and H. Wada. 2005. Complete nucleotide sequence of the mitochondrial genome of Doliolum nationalis with implications for evolution of urochordates. Mol. Phylogenet. Evol. 34: 273–283. Zeng, L.Y. and B.J. Swalla. 2005. Molecular phylogeny of the protochordates: chordate evolution. Can. J. Zool. 83: 24–33. Zrzavý, J. and S. Mihulka, P. Kepka, and A. Bezdek. 1998. Phylogeny of the metazoa based on morphological and 18S ribosomal DNA evidence. Cladistics 14: 249–285. Zrzavý, J. and P. Ríha1, L. Piálek, and J. Janouškovec. 2009. Phylogeny of Annelida (Lophotrochozoa): total-evidence analysis of morphology and six genes. BMC Evol. Biol.9: 189.

Note While this work was in press, several studies of metazoan phylogeny were published. Most notably, Regier et al. (2010) investigated arthropod phylogeny using 62 nuclear proteincoding genes and recovered a well-supported Arthropod tree with Pancrustacea (hexapods within crustaceans) and Mandibulata. Hejnol et al. (2009) found support for acoelomorph monophyly, a basal bilaterian placement for Xenoturbella, and placement of Cycliophora as the sister taxon of Entoprocta. Also, Pick and Philippe et al.’s (2010) investigation of basal metazoan phylogeny recovered the following branching order: (P orifera(Ctenophora(Cnidaria(Placozoa, Bilateria)))). Hejnol, A. and M. Obst, A. Stamatakis, M. Ott, G.W. Rouse, G.D. Edgecombe, P. Martinez, J. Baguña, X. Bally, U. Jondelius, M. Wiens, W.E.G. Müller, E. Seaver, W.C. Wheeler, M.Q. Martindale, G. Giribet, and C.W. Dunn. 2009. Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc. Royal Soc. B. 276(1161): 4261–4270. Regier, J.C. and J.W. Shultz, A. Zwick, A. Hussey, B. Ball, R. Wetzer, J.W. Martin, and C.W. Cunningham. 2010. Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences. Nature 463: 1079–1083. Pick, K.S. and H. Philippe, F. Schreiber, D. Erpenbeck, D.J. Jackson, P. Wrede, M. Wiens, A. Alié, B. Morgenstern, M. Manuel, and G. Wörheide. 2010. Improved phylogenomic taxon sampling noticeably affects non-bilaterian relationships. Molecular Biology and Evolution (in press).

© 2011 by Taylor and Francis Group, LLC

Chapter 3

Key Transitions in Animal Evolution: a Mitochondrial DNA Perspective Dennis V. Lavrov

Introduction When the first complete mitochondrial DNA (mtDNA) sequence—that of humans—was determined (Anderson et al. 1981), it was fittingly described by the phrase “small is beautiful” (Borst and Grivell 1981). Mitochondrial genomes of bilaterian animals are indeed small (~16 kpb), not only because of their limited coding capacity (typically 37 genes), but also due to the remarkable economy of their genomic organization (Fig. 1). Genes encoded in bilaterian mtDNA are compactly arrayed, separated by no, or only a few, nucleotides and typically contain neither introns (but see Valles et al. 2008) nor regulatory sequences. Protein and transfer RNA genes are even often truncated and completed by either posttranscriptional polyadenylation (Yokobori and Pääbo 1997) or, in some cases, editing (Lavrov et al. 2000). Furthermore, changes in the genetic code allowed animals to reduce the set of mitochondrial tRNA genes to 22, several tRNAs fewer than the minimum number required for translation under the standard genetic code (Marck and Grosjean 2002). Encoded ribosomal RNAs are also reduced in size, lacking many secondary structures present in homologous molecules in other groups. In addition to small size, bilaterian mtDNA displays several unusual genetic and genomic 253 Bessey Hall, Department of Ecology, Evolution and Organismal Biology, Iowa State University, Ames, Iowa 50011. E-mail: [email protected]

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

35

features such as unorthodox translation initiation codons, highly modified structures of encoded transfer RNAs, a high rate of sequence evolution, a relatively low rate of gene rearrangements, and the presence of a single large non-coding “control” region (Wolstenholme 1992b). Although the organization of mtDNA is remarkably uniform across different groups of bilaterian animals [but see Armstrong et al. (2000), Helfenbein et al. (2004), and Suga et al. (2008) for some exceptions], it is far from typical for other eukaryotic lineages. In fact, it was known early on that mitochondrial genomes vary greatly in size, gene content, and genome architecture across eukaryotic groups (Wallace 1982, Lang et al. 1999). Interestingly, even the mitochondrial genome of the choanoflagellate Monosiga brevicollis, a close relative of animals, is severalfold larger than “typical” animal mtDNA and harbors 1.5 times as many genes (Fig. 1). It also encodes bacteria-like transfer and ribosomal RNAs, and uses a minimally-derived genetic code with TGA(Trp) as the only deviation from the standard code (Burger et al. 2003). Thus major changes

Fig. 1. mtDNA organization in bilaterian animals (represented by Homo sapiens) and the choanoflagellate Monosiga brevicollis. Protein and ribosomal genes are atp6, atp8–9: subunits 6, 8 and 9 of F0 adenosine triphosphatase (ATP) synthase; cob: apocytochrome b; cox1-3: cytochrome c oxidase subunits 1–3; nad1–6 and nad4L: NADH dehydrogenase subunits 1–6 and 4L; rns and rnl: small and large subunit rRNAs; rps3–19 and rpl2–16: small and large subunit ribosomal proteins; tatC: twin-arginine translocase component C. tRNA genes are identified by the one-letter code for their corresponding amino acid. The drawings are proportional to genome size: M. brevicollis (76,568 bp) and H. sapiens (16,571 bp).

© 2011 by Taylor and Francis Group, LLC

36

Key Transitions in Animal Evolution

in mitochondrial genome architecture co-occurred with the evolution of multicellular animals. Studies of mtDNA in non-bilaterian animals can provide valuable insights into the progression of these changes and also supply data for phylogenetic analyses of animal relationships. Here I review recent progress in our understanding of non-bilaterian mtDNA and discuss the advantages and limitations of mitochondrial datasets for evolutionary and phylogenetic inferences. The mitochondrial DNA of non-bilaterian animals Complete, or nearly-complete, mitochondrial DNA sequences have been determined for >60 species of non-bilaterian animals, including 30 species of cnidarians, 27 species of sponges, and four placozoan strains (Fig. 2, Table 1). These mitochondrial genomes show many deviations from the typical bilaterian mtDNA described above. First, mtDNA is usually larger in size in non-bilaterian compared to bilaterian animals, averaging ~18.0 kbp in Cnidaria, ~20.0 in demosponges, and ~37.4 in placozoans. The larger size of non-bilaterian mtDNA is primarily due to the presence of larger non-coding intergenic regions, as well as more bacteria-like ribosomal and transfer RNA genes (Beagley et al. 1995, Signorovitch et al. 2007, Wang and Lavrov 2008). Second, mtDNA of non-bilaterian animals shows more variation in the gene content. Extra protein-coding genes are

Fig. 2. mtDNA evolution in the Metazoa. Phylogenetic relationships among bilaterian and non-bilaterian animals are currently unresolved and are represented here by a polytomy (empty box). The present review is mostly limited to mtDNA in “non-bilaterian” animals (light-gray box). Only the taxa for which mtDNA data are available are shown. The numbers under the taxon names indicates the number of complete mitochondrial genomes available for each group.

© 2011 by Taylor and Francis Group, LLC

Table 1. Mitochondrial genomes of non-bilaterian animals. Species

GenBank #

Reference

Cnidaria

Species

GenBank

Reference

Demospongiae

Acropora tenuis

NC_003522

(van Oppen et al. 2002)

Agelas schmidti

EU237475

(Lavrov et al. 2008)

Agaricia humilis

NC_008160

(Medina et al. 2006)

Amphimedon compressa

NC_010201

(Erpenbeck et al. 2007) (Lavrov et al. 2008)

Anacropora matthai

NC_006898

None

Amphimedon queenslandica

NC_008944

(Erpenbeck et al. 2007)

Astrangia sp. JVK-2006

NC_008161

(Medina et al. 2006)

Aplysina fulva

NC_010203

(Lavrov et al. 2008)

Aurelia aurita

NC_008446

(Shao et al. 2006)

Axinella corrugata

NC_006894

(Lavrov and Lang 2005b)

Briareum asbestinum

NC_008073

(Medina et al. 2006)

Chondrilla aff. nucula

NC_010208

(Lavrov et al. 2008)

NC_010206

(Lavrov et al. 2008)

(Brugler and France 2007)

Colpophyllia natans

NC_008162

(Medina et al. 2006)

Cinachyrella kuekenthali

EU237479

(Lavrov et al. 2008)

Discosoma sp. CASIZ 168915

NC_008071

(Medina et al. 2006)

Ectyoplasia ferox

EU237480

(Lavrov et al. 2008)

Discosoma sp. CASIZ 168916

NC_008072

(Medina et al. 2006)

Ephydatia muelleri

NC_010202

(Lavrov et al. 2008)

Hydra oligactis

EU237491

(Kayal and Lavrov 2008)

Geodia neptuni

NC_006990

(Lavrov et al. 2005)

Metridium senile

NC_000933

(Beagley et al. 1998)

Halisarca dujardini

NC_010212

(Lavrov et al. 2008)

Montastraea annularis

NC_007224

(Fukami and Knowlton 2005)

Hippospongia lachne

NC_010215

(Lavrov et al. 2008)

Montastraea franksi

NC_007225

(Fukami and Knowlton 2005)

Igernella notabilis

NC_010216

(Lavrov et al. 2008) Table 1 contd...

© 2011 by Taylor and Francis Group, LLC

37

NC_008411

Callyspongia plicifera

Key Transitions in Animal Evolution

Chrysopathes formosa

38

Table 1 contd... NC_007226

(Fukami and Knowlton 2005)

Iotrochota birotulata

NC_010207

(Lavrov et al. 2008)

Montipora cactus

NC_006902

None

Negombata magnifica

NC_010171

(Belinky et al. 2008)

Mussa angulosa

NC_008163

(Medina et al. 2006)

Oscarella carmela

NC_009090

(Wang and Lavrov 2007)

NC_010217

(Lavrov et al. 2008) (Lavrov et al. 2008)

Nematostella sp. JVK2006

NC_008164

(Medina et al. 2006)

Plakortis angulospiculatus

Pavona clavus

NC_008165

(Medina et al. 2006)

Ptilocaulis walpersi

EU237488

Pocillopora eydouxi

NC_009798

(Flot and Tillier 2007)

Suberites domuncula

NC_010496

Pocillopora damicornis

NC_009797

(Flot and Tillier 2007)

Tethya actinia

NC_006991

(Lavrov et al. 2005)

Porites porites

NC_008166

(Medina et al. 2006)

Topsentia ophiraphidites

NC_010204

(Lavrov et al. 2008)

Pseudopterogorgia bipinnata

NC_008157

(Medina et al. 2006)

Vaceletia sp.

NC_010218

(Lavrov et al. 2008)

Rhodactis sp. CASIZ 171755

NC_008158

(Medina et al. 2006)

Xestospongia muta

NC_010211

(Lavrov et al. 2008)

Ricordea florida

NC_008159

(Medina et al. 2006)

Sarcophyton glaucum

AF064823, AF063191

(Beaton et al. 1998)

Hexactinellida

Savalia savaglia

NC_008827

(Sinniger et al. 2007)

Aphrocallistes vastus,

EU000309

(Rosengarten et al., 2008)

Seriatopora caliendrum

NC_010245

(Chen et al. 2008)

Iphiteon panicea

>19,045

(Haen et al. 2007)

Sympagella nux

>16,293

(Haen et al. 2007)

Seriatopora hystrix

NC_010244

(Chen et al. 2008)

Siderastrea radians

NC_008167

(Medina et al. 2006)

NC_008151

(Dellaporta et al. 2006)

Placozoa Trichoplax adhaerens

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

Montastraea faveolata

Species

GenBank #

Reference

BZ49

NC_008833

(Signorovitch et al. 2007)

BZ10101

NC_008832

(Signorovitch et al. 2007)

BZ2423

NC_008834

(Signorovitch et al. 2007)

Species

GenBank

Reference

Key Transitions in Animal Evolution

39

© 2011 by Taylor and Francis Group, LLC

40

Key Transitions in Animal Evolution

found in several lineages, including atp9 for ATP synthase subunit 9 in most demosponges (Wang and Lavrov 2008) and glass sponges (Haen et al. 2007, Rosengarten et al. 2008), tatC for twin-arginine translocase subunit C in Oscarella carmela (Wang and Lavrov 2007), mutS for a putative mismatch repair protein in octocorals (Pont-Kingdon et al. 1995, 1998, Medina et al. 2006), and polB for the DNA-dependent DNA polymerase in the jellyfish Aurelia aurita and one strain of placozoans (Shao et al. 2006, Signorovitch et al. 2007). In addition, out of 22 tRNA genes typically present in mtDNA of bilaterian animals, at most two, trnM and trnW, are found in cnidarians (Beagley et al. 1995, Kayal and Lavrov 2008) and the G1 clade of demosponges (Wang and Lavrov 2008), only five are found in the demosponge Plakortis angulospiculatus (Wang and Lavrov 2008) and 17 are found in another demosponge Amphimedon compressa (Erpenbeck et al. 2007).1 By contrast, additional mitochondrial tRNA genes are present in mtDNA of other demosponges and placozoans. Third, mitochondrial group I introns, with or without the homing endonucleases of the LAGLIDADG type, are found in hexacorals (Beagley et al. 1996, van Oppen et al. 2002, Fukami and Knowlton 2005), placozoans (Signorovitch et al. 2007), and several species of demosponges (Rot et al. 2006, Wang and Lavrov 2008). Fourth, demosponges, cnidarians and placozoans use a minimally-modified genetic code for mitochondrial translation, with TGA=tryptophan as the only deviation. The same genetic code is used for mitochondrial translation in Monosiga brevicollis and in most (but not all) fungi, whereas most bilaterian animals have modified the specificity of at least ATA and AGR codons (Knight et al. 2001). Finally, many nonbilaterian animals display low rates of mitochondrial sequence evolution as revealed by intraspecific (Shearer et al. 2002, Duran et al. 2004, Hellberg 2006, Wörheide 2006) and interspecific (Lavrov et al. 2005, Medina et al. 2006) studies based on mtDNA sequences. A Parallel Evolution of “Bilaterian-Like” mtDNA Remarkably, one group of non-bilaterian animals—Hexactinellida, or glass sponges—evolved mitochondrial genomes very similar to those found in bilaterian animals. Mitochondrial genomes in glass sponges (Hexactinellida) and bilaterian animals share several characteristics, including a nearly identical gene content (37 genes, but with atp9 instead of atp8 in glass sponges), nucleotide composition of the coding strand, and the presence of a single large non-coding region (Haen et al. 2007, Rosengarten et al. 2008). Like those in bilaterian animals, glass sponge mt1 Rosengarten et al. (Rosengarten et al. 2008) claim that three tRNA genes are also missing in the mitochondrial genome of the glass sponge Aphrocallistes vastus. However, we were able to identify the missing tRNAs in the non-coding regions of this genome.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

41

tRNAs display a great degree of variation both in size and in nucleotide sequence of the DHU and TψC arms, including a highly variable sequence of the TψC loop and the lack of conserved guanine residues in the DHU loops. In addition, both groups share a reassignment of the mitochondrial AGR codons from arginine to serine and experienced similar changes in the secondary structure of the tRNASer UCU that translates these codons (Haen et al. 2007). Many of these changes are rare and complex events that, so far, are known to occur only in the Bilateria and Hexactinellida. However, a closer look at mitochondrial genomes from glass sponges and bilaterian animals also reveals some differences. First, although only one mitochondrial gene for a tRNA with the CAU anticodon has been found in both groups, the encoded tRNA in glass sponges does not contain the R11-Y24 pair, characteristic of initiator tRNAs in other organisms. Hence, it is unclear whether this gene codes for the initiator tRNAfMet CAU or for the tRNAIle LAU that usually translates the isoleucine ATA codon in eubacteria and eubacteria-derived organelles, including mitochondria of demosponges and placozoans (see below). If the latter is the case, then glass sponges differ from several groups of bilaterian animals (hemichordates, echinoderms, and flat worms) where ATA also codes for isoleucine, but is translated by a modified tRNAIle GAU together with two other isoleucine codons, AUC and AUU, while the only encoded tRNA with CAU anticodon is clearly an initiator tRNA (Jacobs et al. 1988, Castresana et al. 1998). Second, an atypical R11-Y24 pair is present in tRNAPro UGG/CGG of glass sponges as well as all demosponges and placozoans but is not found in the outgroup species Monosiga brevicollis and Amoebidium parasiticum or in bilaterian animals. Because the R11-Y24 base pair is an important recognition element for the initiator tRNA, it is usually strongly counterselected in elongator tRNAs (Marck and Grosjean 2002), and its presence in the proline tRNA of sponges and T. adhaerens spports the monophyly of non-bilaterian animals or at least the monophyly of the Porifera and Placozoa (this character is not available for Cnidaria, because they lack this and most other tRNA genes in their mtDNA). Finally, many genetic novelties of bilaterian animals, such as unusual initiation codons and incomplete termination codons, are rare or not found in glass sponges. These observations, together with phylogenetic inference based on mitochondrial coding sequences, indicate that shared mitochondrial features between glass sponges and bilaterian animals are the result of parallel evolution rather than a common ancestry.

© 2011 by Taylor and Francis Group, LLC

42

Key Transitions in Animal Evolution

Animal Mitochondrial Synapomorphies Given that most of the traits previously thought to be characteristic of animal mtDNA now appear to have evolved within the Metazoa, one may wonder whether there are any derived mitochondrial features shared by all animals. Our analyses reveal two such putative mitochondrial synapomorphies. First, we found a set of Metazoa-specific insertions/ deletions (indels) in protein-coding genes (Fig. 3) that are well conserved across the Metazoa, but are absent in M. brevicollis and other non-animal species (Lavrov et al. 2005). Although the value of an individual indel for phylogenetic reconstruction is limited (Gribaldo and Philippe 2002), the presence of several such events in mitochondrial genes provides strong support for the monophyly of the Metazoa and can serve as a good indicator of metazoan affinity. Second, an atypical R11-Y24 pair is present in mitochondrial tRNATrp UCA of both bilaterian (Wolstenholme 1992a) and non-bilaterian animals, but is absent in non-metazoan outgroups. As explained above, the R11-Y24 pair is usually strongly counterselected in elongator tRNAs (Marck and Grosjean 2002). The lack of both of these synapomorphies in the mtDNA of M. brevicollis refutes the idea that choanoflagellates may be derived from sponges or other basal metazoans (e.g. King and Carroll 2001, Maldonado 2004). Phylogenetic Inference Using Mitochondrial Sequences and Gene Orders Mitochondrial genomes provide two primary datasets for phylogenetic inference: gene sequences and gene orders (Bruns et al. 1989, Boore and Brown 1998, Lavrov and Lang 2005a). Mitochondrial sequence data

Fig. 3. Conserved indel events shared among multicellular animals. The four best-conserved genes (cob, cox1-3) were analyzed. Numbers above the alignment indicate positions in H. sapiens sequence.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

43

are well suited for phylogenetic analysis within class-level lineages of non-bilaterian animals because of the low rate of sequence evolution and homogeneous nucleotide composition in most of them (Medina et al. 2006, Signorovitch et al. 2007, Wang and Lavrov 2007, Lavrov et al. 2008). For example, the phylogenetic analysis presented in Fig. 4 recovers phylogenetic relationships within the Demospongiae highly congruent with those based on 18S rRNA data and multiple nuclear proteins (see Lavrov et al. 2008 for details). The topology within Hexacorallia also presents a viable phylogenetic hypothesis, largely consistent with previous studies (Berntson et al. 1999, Daly et al. 2003, Collins et al. 2006). Notably, the use of an expanded dataset of anthozoan sequences helps to recover the monophyletic Scleractinia, forming a sister group to naked corals (order Corallimorpharia). This result contradicts the finding of the previous mtDNA-based study (Medina et al. 2006), which inferred the origin of the naked corals within the Scleractinia, but is largely congruent with analyses of other molecular and morphological datasets. At the same time, the inference of global animal relationships based on mtDNA sequences is more problematic. The position of glass sponges is one example. The use of traditional empirical models of amino-acid evolution in maxium likelihood and Bayesian phylogenetic analyses results in a strong support (with high bootstrap and posterior probability numbers) for the sister group relationship between glass sponges and bilaterian animals (Haen et al. 2007). By contrast, Bayesian phylogenetic inference based on a CAT model that explicitly handles the heterogeneity of the substitution process across amino-acid positions (Lartillot and Philippe 2004) places glass sponges either with some fast-evolving cnidarians (Fig. 4) or within demosponges (Haen et al. 2007, Wang and Lavrov 2007). Another recurring result of studies based on mitochondrial protein sequences is the grouping of demosponges with some or all cnidarians (Lavrov et al. 2005, Shao et al. 2006, Haen et al. 2007, Kayal and Lavrov 2008, Lavrov et al. 2008). Although, this grouping may reflect a genuine phylogenetic signal [e.g. see Dunn et al. (2008) for a recent congruent result based on an alternative dataset], it can also be interpreted as an artifact of long-branch attraction (Felsenstein 1978, Hendy and Penny 1989) between rapidly evolving bilaterian animals and distantly-related outgroups (Lavrov et al. 2005). Similar uncertainty surrounds the interpretation of the phylogenetic position of Placozoa, which varies considerably in different analyses and is unresolved in the analysis conducted for this study (Fig. 4). Although one may argue that the missing mitochondrial data from Calcarea and Ctenophora will help to resolve the basal animal relationships, I think that the very absence of their mitochondrial sequences from public databases in this genomic age is symptomatic and indicates highly unusual and fast-evolving mitochondrial genomes in these two groups that will be little informative for phylogenetic reconstruction.

© 2011 by Taylor and Francis Group, LLC

44

Key Transitions in Animal Evolution

Fig. 4. Phylogenetic analysis of global animal relationships using mitochondrial sequence data. Posterior majority-rule consensus tree obtained from the analysis of 2,539 aligned amino acid positions under the CAT+F+Γ model in the PhyloBayes program is shown. We ran four independent chains for ~60,000 generations and sampled every 10th tree after the first 1000 burnin cycles. The convergence among the chains was monitored with the maxdiff statistics and the analysis was terminated after maxdiff became less than 0.3. The number/ circle at each node represents the Bayesian posterior probability. Posterior probability = 1 is indicated by a filled circle; that > 0.95—by an open circle; smaller posterior probabilities are shown as numbers. Amino acid sequences for non-bilaterian animals were derived from the GenBank files listed in table 1. Amino-acid sequences for Cantharellus cibarius mtDNA were downloaded from http://megasun.bch.umontreal.ca/People/lang/FMGP/proteins. html; those for Capsaspora owczarzaki mtDNA were provided by Franz Lang (Université de Montréal). Other sequences were derived from the GenBank files: Arbacia lixula NC_001770, Balanoglossus carnosus NC_001887, Branchiostoma floridae NC_000834, Drosophila yakuba NC_001322, Florometra serratissima NC_001878, Homo sapiens NC_001807, Katharina tunicata NC_001636, Limulus polyphemus NC_003057, Loligo bleekeri NC_002507, Lumbricus terrestris NC_001673, Platynereis dumerilii NC_000931, Priapulus caudatus NC_008557, Saccoglossus kowalevskii NC_007438, Xenoturbella bocki NC_008556, Amoebidium parasiticum AF538042AF538052, Monosiga brevicollis NC_004309, Allomyces macrogynus NC_001715, Mortierella verticillata NC_006838, Rhizopus oryzae NC_006836. Color image of this figure appears in the color plate section at the end of the book.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

45

While gene order comparisons commonly accompany descriptions of new mitochondrial genomes, a formal phylogenetic analyses on this dataset is rarely done. This is mainly due to the lack of user-friendly computer programs for such analyses but also to the algorithmic difficulties in analyzing gene order data (Moret et al. 2004). The paucity of gene order studies is rather unfortunate because they can provide important insights in cases where most other datasets fail, as has been shown on echinoderms (Smith et. al 1993), arthropods (Boore et. al 1995, Boore et al. 1998), and crustaceans (Lavrov et al. 2004, Morrison et al. 2002). Recently we used mitochondrial gene arrangements to infer phylogenetic relationships within the Demospongiae and recovered a tree largely congruent with that based on mitochondrial sequences, although with only a moderate support (Lavrov et al. 2008). However, the analysis of global animal phylogeny based on mitochondrial gene order data is again problematic and results in a phylogeny were most of the basal relationships are unresolved (Fig. 5). We note that there are several problems complicating mitochondrial gene-order based analysis in non-bilaterian animals. These include the lack of informative data from accepted outgroups, variation in the mitochondrial gene content across non-bilaterian animals, especially, an independent loss of mitochondrial tRNA genes in several lineages, and an unusual mitochondrial genomic organization in hexacorrals and placozoans, where some mitochondrial genes are located within introns of other genes. Mitochondrial Size and Gene Content are Not Reliable Phylogenetic Indicators Can mitochondrial genome size and gene content be used for phylogenetic reconstruction? At first glance there appears to be a trend in animal mtDNA evolution, with both genome size and gene content being reduced during transitions first to the Metazoa and then to the Bilateria (Fig. 2). Indeed, the mitochondrial genome of the choanoflagellate Monosiga brevicollis is 76.6 kbp in size and contains 55 genes, those of most demosponges are between 18 and 25 kbp and contain 40–44 genes, while bilaterian mtDNA is typically between 14 and 16 kpb and has 36–37 genes. Thus, it may be tempting to use mtDNA size and gene content as indicators of phylogenetic relationships (e.g. Dellaporta et al. 2006). It should be noted, however, that the reduction in mitochondrial DNA size and gene content are only overall trends in animal mitochondrial evolution and that both of these features can vary substantially within individual groups. The largest animal mitochondrial genomes (up to 43 kbp) are found in the placozoan Trichoplax adhaerens (Dellaporta et al. 2006) as well as some very distantly related bilaterian animals, including three species of

© 2011 by Taylor and Francis Group, LLC

46

Key Transitions in Animal Evolution

bark weevil Pissodes (Boyce et al. 1989), the deep-sea scallop Placopecten magellanicus (Snyder et al. 1987), and the nematode Romanomermis culicivorax (Powers et al. 1986), and have clearly evolved independently in these taxa. In addition, relatively closely related animals often display extensive variation in mtDNA size. For example, the size of the mitochondrial genome in Drosophila melanogaster is ~22% (3.5 kpb) larger than that in Drosophila yakuba, while the mtDNA length in four strains of Placozoa differs by as much as 35% (Signorovitch et al. 2007). The fact that mtDNA size can change rapidly and repeatedly within independent lineages makes it an unreliable character for phylogenetic reconstruction. Furthermore, it is likely that we underestimate the true range of length variation in bilaterian mtDNA because of the difficulties associated with PCR amplification and sequencing of the large non-coding region in this molecule (e.g. Lavrov and Brown 2001).

Fig. 5. Global animal relationships based on mitochondrial gene order data. Strict consensus tree is shown from the Maximum Parsimony analysis on Multistate Encodings (Boore et al. 1995, Wang et al. 2002). Gene orders were encoded as described previously (Lavrov and Lang 2005a) and analyzed using heuristic search with 100 random addition replicates in PAUP*4.0b10 and TBR branch swapping option. Color image of this figure appears in the color plate section at the end of the book.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

47

Mitochondrial gene content is also a poor indicator of phylogenetic relationships. In most cases when “extra” genes are present, they are inherited from a common ancestor and thus represent plesiomorphies not informative for phylogenetic reconstruction. By contrast, while the losses of genes from mtDNA are apomorphies, they are known to occur in parallel in different groups, resulting in homoplasious similarities in gene contents (Martin et al. 1998). As an example, a very similar mitochondrial gene content in animal and fungal mtDNA has likely evolved independently in the two groups given that the mitochondrial genome of the choanoflagellate Monosiga brevicollis, the sister group to animals, contains additional genes (Burger et al. 2003; Fig. 1). Another example comes from our recent study of mitochondrial genomes in demosponges. This study revealed that the G1 group of demosponges lost all but two tRNA genes from mtDNA, resulting in a gene content very similar to that in the phylum Cnidaria. Remarkably, in both of these groups the same two tRNA genes have been retained (trnM(cau) and trnW(uca)) supporting the inference of a special role of these tRNAs in animal mitochondria. As has been explained before (e.g. Shao et

tRNAMet CAU is used for the initiation of mitochondrial translation Trp with formyl-methionine (Smith and Marcker 1968) while tRNAUCA must al. 2006),

translate the TGA in addition to the TGG codons as tryptophan in animal mitochondria. Because of these special functions, the transfer of these tRNAs to the nucleus may be selectively disadvantageous. Implications for mtDNA Evolution Although mitochondrial genome size and gene content offer little information for resolving animal phylogenetic relationships, they do provide insights into mitochondrial genome evolution. Given that the transfer of nuclear genes to mitochondria is limited (in part by differences in genetic code), most additional genes found in demosponge mitochondrial DNA (atp9, tatC, trnI(cau), trnR(ucu)) were likely inherited from their common ancestor with animals. This is particularly the case for trnI(cau). The maturation of tRNAIle CAU encoded by this gene involves a posttranscriptional modification of the cytosine at position 34 to lysidine (2-lysyl-cytidine) (Muramatsu et al. 1988, Weber et al. 1990). This modification is performed by the tRNAIle-lysidine synthetase (Soma et al. 2003), an enzyme that is not involved in the maturation of cytoplasmic isoleucine tRNAs (Marck and Grosjean 2002). Because of the dispensable nature of this function for cytoplasmic protein synthesis, it is likely that in Ile evolutionary terms, the loss of the metazoan mitochondrial tRNACAU is followed by the loss of the nuclear lysidine synthetase gene, rendering a reacquisition of tRNAIle practically impossible. The presence of trnI(cau) CAU

© 2011 by Taylor and Francis Group, LLC

48

Key Transitions in Animal Evolution

in the mtDNA of both demosponges and placozoans clearly indicates that the common animal ancestor used a less modified genetic code for mitochondrial translation than do most bilaterian animals. It is also likely that this common ancestor had more genes in its mitochondrial genome than extant metazoan taxa. Signorovitch et al. (2007) suggest that the common ancestor of all animals possessed a large, noncompact mitochondrial genome. This inference is based on the observation that both placozoans and the choanoflagellate Monosiga brevicollis have large mtDNA and on the assumption that Placozoa forms the sister group to all other animals. There are two potential problems with this inference. First, phylogenetic analysis of global animal relationships based on mtDNA sequence data does not support the placement of Placozoa as the sister group to other animals (Signorovitch et al. 2007, Wang and Lavrov 2007). Second, even if Placozoa is the sister group to the rest of the animals, the most parsimonious reconstruction of genome size in the common animal ancestor can be grossly misleading (e.g. Cunningham et al. 1998) given the long branch leading to the Placozoa and rapid changes in the size of mtDNA observed among closely-related animals (see above). A more interesting question, in my view, is about the evolutionary forces that maintain the compact nature of mtDNA in most modern animals. Studies of non-bilaterian animals can help to answer this question because these animals display a different combination of features in their mitochondrial organization. For example, these studies indicate that small size and gene content are not directly linked with an accelerated rate of evolution of mitochondrial sequences. As shown above, while the rate of nucleotide substitutions appears to be extremely low in some nonbilaterian animals, their genomes are still very compact and mostly intronless (with the exception of Placozoa), challenging the idea that elevated mutation pressure is solely responsible for the evolution of these features in animal mtDNA (Lynch et al. 2006). Implications for Animal Evolution It is commonly believed that the study of non-bilaterian animals can also provide insights into metazoan morphological evolution. This is not because non-bilaterian animals are phylogenetically more “basal” or “lower”—these terms are misleading and should be avoided (Crisp and Cook 2005)—but because they evolved their bodyplans earlier in metazoan evolution (as revealed by the fossil record) and so may resemble the ancestors of other groups. Following this reasoning, sponges are often considered as living representatives of an intermediate stage in animal evolution that never reached the tissue-level grade of organization. Our

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

49

recent study of mtDNA from the homoscleromorph Oscarella carmela challenges this idea (Wang and Lavrov 2007). Homoscleromorpha is a small group of sponges that share? several features with “higher” animals such as the presence of type-IV collagen, acrosomes in spermatozoa and cross-striated rootlets in the flagellar basal apparatus of larval cells. More importantly, it has been shown that the epithelial cells in homoscleromorph larvae meet all the criteria of a true epithelium in higher animals: cell polarization, apical cell junctions, and a basement membrane (Boury-Esnault et al. 2003). Unless these shared cytological features evolved independently in Homoscleromorpha and Eumetazoa (an unlikely scenario), two alternative explanations are possible for these findings: either Homoscleromorpha is more closely related to other animals than to sponges or most sponges lost the aforementioned features. Our analysis of demosponge relationships based on mitochondrial genomic data did not find any support for the first of these hypotheses and instead provided strong support for the placement of the Homoscleromorpha with demosponges. This result suggests that the bodyplan of sponges might represent a secondary simplification in animal morphology, potentially due to their sedentary and water-filtering lifestyle. Similar conclusion should be made if sponges and cnidarians form a monophyletic group (Fig. 1) or if ctenophores form the sister group to the rest of the animals (Dunn et al. 2008). Conclusions The evolution of multicellular animals is associated with major changes in their mitochondrial genome architecture. Recent studies on mtDNA from non-bilaterian animals suggest that these changes occurred in two steps, roughly correlated with the origin of animal multicellularity and the origin of the bilaterality. The transition to multicellular animals is associated with the loss of multiple genes from mtDNA and a drastic reduction in the amount of non-coding DNA in the genome, resulting in its “small is beautiful” nature. The transition to bilaterian animals is correlated with multiple changes in the genetic code and associated losses of tRNA genes, the emergence of several genetic novelties, and a large increase in the rates of sequence evolution. Although we do not know whether the observed changes co-occurred with the morphological transitions or evolved independently in different lineages, the remarkable uniformity of animal mtDNA suggests an ongoing evolutionary pressure that maintains its unique organization. Studies of mtDNA in non-bilaterian animals are important because they provide insights not only into the history of mtDNA evolution, but also into the evolutionary factors that shape modern-day animal mtDNA. Although our sampling of mtDNA from non-bilaterian animals is still limited, substantial progress has been made in the last few

© 2011 by Taylor and Francis Group, LLC

50

Key Transitions in Animal Evolution

years that revealed new information about phylogenetics, the evolution of mitrochondrial genomes, and even morphological evolution in animals. Acknowledgements I thank Bernd Schierwater, Stephen Dellaporta, and Rob DeSalle for organizing the symposium and its complementary session, Karri Haen, Ehsan Kayal and Xiujuan Wang for valuable comments on an earlier version of this manuscript, and the College of Liberal Arts and Sciences at Iowa State University for funding. References Anderson, S. and A.T. Bankier, B.G. Barrell, M.H.L. De Bruijn, A.R. Coulson, J. Drouin et al. 1981. Sequence, and organization of the human mitochondrial genome. Nature 290: 457–465. Armstrong, M.R. and V.C. Block, and M.S. Phillips. 2000. A multipartite mitochondrial genome in the potato cyst nematode Globodera pallida. Genetics 154: 181–192. Beagley, C.T. and J.L. Macfarlane, G.A. Pont-Kingdon, R. Okimoto, N. Okada, and D.R. Wolstenholme. 1995. Mitochondrial genomes of Anthozoa (Cnidaria). Palmieri F., Papa S., Saccone C., and Gadaleta N. (eds) Progress in cell research – Symposium on “Thirty Years of Progress in Mitochondrial Bioenergetics and Molecular Biology”. Elsevier Science BV Amsterdam, The Netherlands, pp 149–153. Beagley, C.T. and N.A. Okada, and D.R. Wolstenholme. 1996. Two mitochondrial group I introns in a metazoan, the sea anemone Metridium senile: one intron contains genes for subunits 1 and 3 of NADH dehydrogenase. Proc. Natl. Acad. Sci. USA 93: 5619–5623. Beagley, C.T. and R. Okimoto, and D.R. Wolstenholme. 1998. The mitochondrial genome of the sea anemone Metridium senile (Cnidaria): introns, a paucity of tRNA genes, and a near-standard genetic code. Genetics 148: 1091–1108. Beaton, M.J. and A.J. Roger, and T. Cavalier-Smith. 1998. Sequence analysis of the mitochondrial genome of Sarcophyton glaucum: conserved gene order among octocorals. J. Mol. Evol. 47: 697–708. Belinky, F. and C. Rot, M. Ilan and D. Huchon. 2008. The complete mitochondrial genome of the demosponge Negombata magnifica (Poecilosclerida). Mol. Phylogenet. Evol. 47: 1238–1243. Berntson, E.A. and S.C. France, and L.S. Mullineaux. 1999. Phylogenetic relationships within the class Anthozoa (phylum Cnidaria) based on nuclear 18S rDNA sequences. Mol. Phylogenet. Evol. 13: 417–433. Boore, J.L. and T.M. Collins, D. Stanton, L.L. Daehler, and W.M. Brown. 1995. Deducing the pattern of arthropod phylogeny from mitochondrial DNA rearrangements. Nature 376: 163–165. Boore, J.L. and W.M. Brown. 1998. Big trees from little genomes: mitochondrial gene order as a phylogenetic tool. Curr. Opin. Genet. Dev. 8: 668–674. Borst, P. and L.A. Grivell. 1981. Small is beautiful—portrait of a mitochondrial genome. Nature 290: 443–444. Boury-Esnault, N. and A. Ereskovsky, C. Bezac, and D. Tokina. 2003. Larval development in the Homoscleromorpha (Porifera, Demospongiae). Invertebr. Biol. 122: 187–202. Boyce, T. M. and M.E. Zwick, and C.F. Aquadro. 1989. Mitochondrial DNA in the bark weevils: Size, structure and heteroplasmy. Genetics 123: 825–836. Brugler, M.R. and S.C. France. 2007. The complete mitochondrial genome of the black coral Chrysopathes formosa (Cnidaria:Anthozoa:Antipatharia) supports classification of antipatharians within the subclass Hexacorallia. Mol. Phylogenet. Evol. 42: 776–788.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

51

Bruns, T.D. and R. Fogel, T.J. White, and J.D. Palmer. 1989. Accelerated evolution of a falsetruffle from a mushroom ancestor. Nature 339: 140–142. Burger, G. and L. Forget, Y. Zhu, M.W. Gray, and B.F. Lang. 2003. Unique mitochondrial genome architecture in unicellular relatives of animals. Proc. Natl. Acad. Sci. USA 100: 892–897. Castresana, J. and G. Feldmaier-Fuchs, S. Yokobori, N. Satoh and S. Paabo. 1998. The mitochondrial genome of the hemichordate Balanoglossus carnosus and the evolution of deuterostome mitochondria. Genetics 150: 1115–1123. Chen, C. and C.F. Dai, S. Plathong, C.Y. Chiou, and C.A. Chen. 2008. The complete mitochondrial genomes of needle corals, Seriatopora spp. (Scleractinia: Pocilloporidae): an idiosyncratic atp8, duplicated trnW gene, and hypervariable regions used to determine species phylogenies and recently diverged populations. Mol. Phylogenet. Evol. 46: 19–33. Collins, A.G. and P. Schuchert, A.C. Marques, T. Jankowski, M. Medina, and B. Schierwater. 2006. Medusozoan phylogeny and character evolution clarified by new large and small subunit rDNA data and an assessment of the utility of phylogenetic mixture models. Syst. Biol. 55: 97–115. Crisp, M.D. and L.G. Cook. 2005. Do early branching lineages signify ancestral traits? Trends Ecol. Evol. 20: 122–128. Cunningham, C.W. and K.E. Omland, and T.H. Oakley. 1998. Reconstructing ancestral character states: a critical reappraisal. Trends Ecol. Evol. 13: 361–366. Daly, M. and D.G. Fautin, and V.A. Cappola. 2003. Systematics of the Hexacorallia (Cnidaria : Anthozoa). Zool. J. Linn. Soc. Lond. 139: 419–437. Dellaporta, S.L. and A. Xu, S. Sagasser, W. Jakob, M.A. Moreno, L.W. Buss, and B. Schierwater. 2006. Mitochondrial genome of Trichoplax adhaerens supports Placozoa as the basal lower metazoan phylum. Proc. Natl. Acad. Sci. USA 103: 8751–8756. Dunn, C.W. and A. Hejnol, D.Q. Matus, K. Pang, W.E. Browne, S.A. Smith et al. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–749. Duran, S. and M. Pascual, and X. Turon. 2004. Low levels of genetic variation in mtDNA sequences over the western Mediterranean and Atlantic range of the sponge Crambe crambe (Poecilosclerida). Mar. Biol. 144: 31–35. Erpenbeck, D. and O. Voigt, M. Adamski, M. Adamska, J. Hooper, G. Worheide, and B. Degnan. 2007. Mitochondrial diversity of early-branching Metazoa is revealed by the complete mt genome of a haplosclerid demosponge. Mol. Biol. Evol. 24: 19–22. Felsenstein, J. 1978. Cases in which parsimony or compatibility methods will be positively misleading. Syst. Zool. 27: 401–410. Flot, J.F. and S. Tillier. 2007. The mitochondrial genome of Pocillopora (Cnidaria: Scleractinia) contains two variable regions: the putative D-loop and a novel ORF of unknown function. Gene 401: 80–87. Fukami, H. and N. Knowlton. 2005. Analysis of complete mitochondrial DNA sequences of three members of the Montastraea annularis coral species complex (Cnidaria, Anthozoa, Scleractinia). Coral Reefs 24: 410–417. Gribaldo, S. and H. Philippe. 2002. Ancient phylogenetic relationships. Theor. Popul. Biol. 61: 391–408. Haen, K.M. and B.F. Lang, S.A. Pomponi, and D.V. Lavrov. 2007. Glass sponges and bilaterian animals share derived mitochondrial genomic features: A common ancestry or parallel evolution? Mol. Biol. Evol. 24: 1518–1527. Helfenbein, K.G. and H.M. Fourcade, R.G. Vanjani, and J.L. Boore. 2004. The mitochondrial genome of Paraspadella gotoi is highly reduced and reveals that chaetognaths are a sister group to protostomes. Proc. Natl. Acad. Sci. USA 101: 10639–10643. Hellberg, M.E. 2006. No variation and low synonymous substitution rates in coral mtDNA despite high nuclear variation. BMC Evol. Biol. 6: 24.

© 2011 by Taylor and Francis Group, LLC

52

Key Transitions in Animal Evolution

Hendy, M.D. and D. Penny. 1989. A framework for the quantitative study of evolutionary trees. Syst. Zool. 38: 297–309. Jacobs, H.T. and D.J. Elliott, V.B. Math, and A. Farquharson. 1988. Nucleotide sequence and gene organization of sea urchin mitochondrial DNA. J. Mol. Biol. 202: 185–217. Kayal, E. and D.V. Lavrov. 2008. The mitochondrial genome of Hydra oligactis (Cnidaria, Hydrozoa) sheds new light on animal mtDNA evolution and cnidarian phylogeny. Gene 410: 177–186. King, N. and S.B. Carroll. 2001. A receptor tyrosine kinase from choanoflagellates: molecular insights into early animal evolution. Proc. Natl. Acad. Sci. USA 98: 15032–15037. Knight, R.D. and S.J. Freeland, and L.F. Landweber. 2001. Rewiring the keyboard: evolvability of the genetic code. Nat. Rev. Genet. 2: 49–58. Lang, B.F. and M.W. Gray, and G. Burger. 1999. Mitochondrial genome evolution and the origin of eukaryotes. Annu. Rev. Genet. 33: 351–397. Lartillot, N. and H. Philippe. 2004. A Bayesian mixture model for across-site heterogeneities in the amino-acid replacement process. Mol. Biol. Evol. 21: 1095–1109. Lavrov, D.V. and W.M. Brown, and J.L. Boore. 2000. A novel type of RNA editing occurs in the mitochondrial tRNAs of the centipede Lithobius forficatus. Proc. Natl. Acad. Sci. USA 97: 13738–13742. Lavrov, D.V. and W.M. Brown. 2001. Trichinella spiralis mtDNA: A nematode mitochondrial genome that encodes a putative ATP8, normally-structured tRNAs, and has a gene arrangement relatable to those of coelomate metazoans. Genetics 157: 621–637. Lavrov, D.V. and L. Forget, M. Kelly, and B.F. Lang. 2005. Mitochondrial genomes of two demosponges provide insights into an early stage of animal evolution. Mol. Biol. Evol. 22: 1231–1239. Lavrov, D.V. and B.F. Lang. 2005a. Poriferan mtDNA and animal phylogeny based on mitochondrial gene arrangements. Syst. Biol. 54: 651–659. Lavrov, D.V. and B.F. Lang. 2005b. Transfer RNA gene recruitment in mitochondrial DNA. Trends Genet. 21: 129–133. Lavrov, D.V. and M. Kelly, and X. Wang. 2008. Reconstructing ordinal relationships in the Demospongiae using mitochondrial genomic data. Mol. Phylogenet. Evol. 49: 111–124. Lynch, M. and B. Koskella, and S. Schaack. 2006. Mutation pressure and the evolution of organelle genomic architecture. Science 311: 1727–1730. Maldonado, M. 2004. Choanoflagellates, choanocytes, and animal multicellularity. Invertebr. Biol. 123: 1–22. Marck, C. and H. Grosjean. 2002. tRNomics: analysis of tRNA genes from 50 genomes of Eukarya, Archaea, and Bacteria reveals anticodon-sparing strategies and domainspecific features. RNA 8: 1189–1232. Martin, W. and B. Stoebe, V. Goremykin, S. Hapsmann, M. Hasegawa, and K.V. Kowallik. 1998. Gene transfer to the nucleus and the evolution of chloroplasts. Nature 393: 162–165. Medina, M. and A.G. Collins, T.L. Takaoka, J.V. Kuehl, and J.L. Boore. 2006. Naked corals: skeleton loss in Scleractinia. Proc. Natl. Acad. Sci. USA 103: 9096–9100. Moret, B.M. and J. Tang, and T. Warnow. Reconstructing phylogenies from gene-content and gene-order data. pp. 1–32. In: O. Gascuel. [ed.] 2004. Mathematics of Evolution and Phylogeny. Clarendon Press, Oxford. Muramatsu, T. and K. Nishikawa, F. Nemoto, Y. Kuchino, S. Nishimura, T. Miyazawa, and S. Yokoyama. 1988. Codon and amino-acid specificities of a transfer RNA are both converted by a single post-transcriptional modification. Nature 336: 179–181. Pont-Kingdon, G.A. and N.A. Okada, J.L. Macfarlane, C.T. Beagley, D.R. Wolstenholme, T. Cavalier-Smith, and G.D. Clark-Walker. 1995. A coral mitochondrial mutS gene. Nature 375: 109–111. Pont-Kingdon, G. and N.A. Okada, J.L. Macfarlane, C.T. Beagley, C.D. Watkins-Sims, T. CavalierSmith, G.D. Clark-Walker, and D.R. Wolstenholme. 1998. Mitochondrial DNA of the coral Sarcophyton glaucum contains a gene for a homologue of bacterial MutS: a possible case of gene transfer from the nucleus to the mitochondrion. J. Mol. Evol. 46: 419–431.

© 2011 by Taylor and Francis Group, LLC

Key Transitions in Animal Evolution

53

Powers, T.O. and E.G. Platzer, and B.C. Hyman. 1986. Large mitochondrial genome and mitochondrial DNA size polymorphism in the mosquito parasite, Romanomermis culicivorax. Curr. Genet. 11: 71–77. Rosengarten, R.D. and E.A. Sperling, M.A. Moreno, S.P. Leys, and S.L. Dellaporta. 2008. The mitochondrial genome of the hexactinellid sponge Aphrocallistes vastus: evidence for programmed translational frameshifting. BMC Genomics 9: 33. Rot, C. and I. Goldfarb, M. Ilan, and D. Huchon. 2006. Putative cross-kingdom horizontal gene transfer in sponge (Porifera) mitochondria. BMC Evol. Biol. 6: 71. Shao, Z. and S. Graf, O.Y. Chaga, and D.V. Lavrov. 2006. Mitochondrial genome of the moon jelly Aurelia aurita (Cnidaria, Scyphozoa): A linear DNA molecule encoding a putative DNA-dependent DNA polymerase. Gene 381: 92–101. Shearer, T.L. and M.J. Van Oppen, S.L. Romano, and G. Worheide. 2002. Slow mitochondrial DNA sequence evolution in the Anthozoa (Cnidaria). Mol. Ecol. 11: 2475–2487. Signorovitch, A.Y. and L.W. Buss, and S.L. Dellaporta. 2007. Comparative genomics of large mitochondria in placozoans. PLoS Genet. 3: e13. Sinniger, F. and P. Chevaldonne, and J. Pawlowski. 2007. Mitochondrial genome of Savalia savaglia (Cnidaria, Hexacorallia) and early metazoan phylogeny. J. Mol. Evol. 64: 196–203. Smith, A.E. and K.A. Marcker. 1968. N-formylmethionyl transfer RNA in mitochondria from yeast and rat liver. J. Mol. Biol. 38: 241–243. Snyder, M. and A.R. Fraser, J. Laroche, K.E. Gartner-Kepkay, and E. Zouros. 1987. Atypical mitochondrial DNA from the deep-sea scallop Placopecten magellanicus. Proc. Natl. Acad. Sci. USA 84: 7595–7599. Soma, A. and Y. Ikeuchi, S. Kanemasa, K. Kobayashi, N. Ogasawara, T. Ote, J. Kato, K. Watanabe, Y. Sekine, and T. Suzuki. 2003. An RNA-modifying enzyme that governs both the codon and amino acid specificities of isoleucine tRNA. Mol. Cell. 12: 689–698. Suga, K. and D.B. Mark Welch, Y. Tanaka, Y. Sakakura, and A. Hagiwara. 2008. Two circular chromosomes of unequal copy number make up the mitochondrial genome of the rotifer Brachionus plicatilis. Mol. Biol. Evol. 25: 1129–1137. Valles, Y. and K.M. Halanych, and J.L. Boore. 2008. Group II introns break new boundaries: presence in a bilaterian’s genome. PLoS ONE 3: e1488. van Oppen, M.J. and J. Catmull, B.J. McDonald, N.R. Hislop, P.J. Hagerman, and D.J. Miller. 2002. The mitochondrial genome of Acropora tenuis (Cnidaria; Scleractinia) contains a large group I intron and a candidate control region. J. Mol. Evol. 55: 1–13. Wallace, D.C. 1982. Structure and evolution of organelle genomes. Microbiol. Rev. 46: 208–240. Wang, L.S. and R.K. Jansen, B.M. Moret, L.A. Raubeson, and T. Warnow. 2002. Fast phylogenetic methods for the analysis of genome rearrangement data: an empirical study. Proc. 7th Pacific Symp. on Biocomputing (PSB’02) 524–535. Wang, X. and D.V. Lavrov. 2007. Mitochondrial genome of the homoscleromorph Oscarella carmela (Porifera, Demospongiae) reveals unexpected complexity in the common ancestor of sponges and other animals. Mol. Biol. Evol. 24: 363–373. Wang, X. and D.V. Lavrov. 2008. Seventeen new complete mtDNA sequences reveal extensive mitochondrial genome evolution within the Demospongiae. PLoS ONE 3: e2723. Weber, F. and A. Dietrich, J.H. Weil, and L. Maréchal-Drouard. 1990. A potato mitochondrial isoleucine tRNA is coded for by a mitochondrial gene possessing a methionine anticodon. Nucleic Acids Res. 18: 5027–5030. Wolstenholme, D.R. 1992a. Animal mitochondrial DNA: structure and evolution. Int. Rev. Cytol. 141: 173–216. Wolstenholme, D.R. 1992b. Genetic novelties in mitochondrial genomes of multicellular animals. Curr. Opin. Genet. Dev. 2: 918–925. Wörheide, G. 2006. Low variation in partial cytochrome oxidase subunit I (COI) mitochondrial sequences in the coralline demosponge Astrosclera willeyana across the Indo-Pacific. Mar. Biol. 148: 907–912. Yokobori, S. and S. Pääbo. 1997. Polyadenylation creates the discriminator nucleotide of chicken mitochondrial tRNA(Tyr). J. Mol. Biol. 265: 95–99.

© 2011 by Taylor and Francis Group, LLC

Chapter 4

Pending Issues in Development and Phylogeny of Arthropods Jean S. Deutsch

Presentation Arthropoda is by far the most numerous and most speciose phylum of the animal kingdom. From antiquity, arthropods have attracted the interest of naturalists. During the last century, an arthropod, the fly Drosophila melanogaster has been the model animal in many fields of biology, and can be viewed as the ‘source’ of modern genetics (Morgan 1928) and developmental genetics (Lewis 1978; Nüsslein-Volhard and Wieschaus 1980). Still, at the dawn of the 21st century, many questions remain unsolved about the phylogeny of arthropods, which most often are related to development. In the present chapter, due to limited space, I will present only some of the pending issues, this choice being obviously personal and arbitrary. In some cases, in addition of presenting pros and cons, I will give my personal opinion, always debatable. I apologize to those who will not find discussion on their preferred group or topic. Because I will often focus on molecular and on developmental data, I will keep aside the fossil taxa, although I acknowledge the value of paleontological data for phylogeny.

UMR 7622 Biologie du Développement, CNRS et Université P. et M. Curie (Paris 6); case 24, 9 quai St-Bernard, 75252 Paris Cedex 05, France. E-mail: [email protected]

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

55

Homology, Evolution and Development It is far beyond the scope of the present chapter to discuss exhaustively on what is ‘homology’, one of the most important, most difficult and debated concept in evolutionary biology. Hereafter, I will discuss homology between different species on a structural basis, by analysing the relationships between the various elements composing the structure of interest. This is valid at every level of biological organisation, from the whole body plan to organs, tissues, cells, molecules. This approach was first developed by Étienne Geoffroy Saint-Hilaire (1818) as “the principle of connections”; see Scholtz (2005) for an extended and modern view. This structural approach is not contradictory with the Darwinian concept of homology as “similarity that resulted from common descent” (“homogeny” by Lankester 1870). Rather, it provides the best practical criterion for deriving hypotheses on homologous characters (de Beer 1938 p. 70). Indeed, any phylogenetic analysis requires constructing a set (data matrix) of homologous characters between the different taxa under study (primary homology). We all know how much molecular phylogenetic analyses depend on a correct alignment, which is nothing but application of Geoffroy’s principle of connections to molecules. Owen (1849) distinguished “special homology” that we can today equate to homology by descent (Amundson 2007), from “serial homology”, which is homology between parts within the same individual. Darwin’s own definition of homology in the glossary of the 6th edition of the Origin of Species (1872) takes both types into account. Serial homology is not due to common descent, but derives from the same processes (including similar genetic networks) in constructing different organs. The same structural analysis can be used to characterize both types of homologues. The Position of the Arthropoda Within the Animal Kingdom Cuvier (1817) grouped the Arthropoda with the Annelida, into the ‘Articulata’, one of his four ‘Embranchements’ of the animal kingdom. The same idea is reflected in the etymology of the Greek word ‘En-tomon’ and Latin ‘In-secta’, or segmented animals. In 1997, Aguinaldo et al., after the results of a molecular phylogenetic analysis of 18S RNA, coined the name ‘Ecdysozoa’ for a clade of moulting animals—‘ecdysis’ is the Greek word for ‘moult’—grouping the Introverta (Nielsen 2001) (i.e. Nematoda, Nematomorpha, Priapulida, Kinoryncha and Loricifera)1 with the Arthropoda and related phyla, Onychophora (velvet worms) (Ballard et al. 1992) and Tardigrada (water bears) (Giribet 1 The phylogenetic affinities of the Gastrotricha are still much debated. They have been placed either without or within the Ecdysozoa or again as their sister-group. This controversy is out of the scope of the present discussion.

© 2011 by Taylor and Francis Group, LLC

56

Key Transitions in Animal Evolution

et al. 1996). This was the start of the so-called ‘new phylogeny’ (Adoutte et al. 2000) proposing a major revision of the previous phylogenetic picture of the animal kingdom. Although still debated [see Nielsen (2001, pp. 119, 510–519), Brusca and Brusca (2003, p. 499), Scholtz (2002, 2003), Nielsen 2003, and Jenner and Scholtz (2005) for extensive discussion] the Ecdysozoa grouping gained more and more support among the community of zoologists, on the basis of both molecular and morphological (Schmidt-Rhaesa et al. 1998, Peterson and Eernisse 2001) studies. Curiously, to accommodate the ‘Articulata’ grouping, Nielsen (2001, 123–137, 215–217) still includes the Arthropoda within the Spiralia, protostome animals whose embryo develops through a spiral cleavage, thus following Anderson (1973). Evidence for arthropods being spiralians has been discussed by Costello and Henley (1976) and evidence against has been reported by Scholtz (1997). It is worth mentioning the work of de Rosa et al. (1999), who examined Hox genes, developmental genes known to have a critical role in the design of the body plan. The Hox genes’ complement in a variety of bilaterian phyla support the dispatching of the Bilateria into three ‘superphyla’, namely Deuterostomia, Lophotrochozoa and Ecdysozoa. As for the present purpose, the main outcome of the ‘new phylogeny’ is the dismissal of the ‘Articulata’. Monophyly of the Arthropoda: a Story of Appendages ‘Arthropoda’ means ‘articulated legs’. It is thus an important diagnostic character. Monophyly of the Arthropoda has been disputed by Sidnie Manton and co-workers (see Fryer 1996, 1997), on the basis of differences in appendages: mainly branching or not of the limbs and articulation of the mandible. Crustacea, possessing limbs with two branches (biramous), would have been issued from an annelid-like stem by a convergent process of ‘arthropodization’ of a shoot different from that of Onychophora, Hexapoda and Myriapoda, together called ‘Uniramia’. Under the polyphyletic hypothesis Chelicerata would be issued from another branch (Fig. 1A). On the opposite side, homology of the appendages among arthropods has been supported by a structural analysis by Snodgrass (1935, Chap. 5): all arthropod legs are built on a similar pattern of ‘podomeres’ (leg segments). More recently, evo-devo approaches have shown that the construction of legs in all arthropod classes depends on the same network of interacting transcription factors and signalling molecules as the one which has been dissected in detail in Drosophila (Abu-Shaar and Mann 1998): Crustacea (Panganiban et al. 1995, Williams 1998, Nulsen and Nagy 1999, Abzhanov and Kaufman 2000, Williams et al. 2002, Hejnol and Scholtz 2004, Pripc

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

57

Fig. 1. Polyphyly vs. monophyly of the Arthropoda. A. The polyphyletic hypothesis. B. The monophyletic hypothesis.

and Telford 2008, Sewell et al. 2008, Williams 2008); Myriapoda (Prpic and Tautz 2003, Prpic 2004); Chelicerata (Abzhanov and Kaufman 2000, Prpic et al. 2003). There are some variations in the expression of the corresponding genes along the proximal to distal axis (e.g. Prpic et al. 2003, Popadic 2005) but they do not overcome those observed within the Hexapoda (e.g. Jockusch et al. 2000, Jockusch et al. 2004). This common genetic system, including wingless (wg/Wnt-1) (Nulsen and Nagy 1999); Distal-less (Dll) (Panganiban et al. 1995, Williams 1998, Williams et al. 2002, Hejnol and Scholtz 2004, Williams 2008), extradenticle (exd) (Williams et al. 2002) and nubbin (nub) (Averof and Cohen 1997, Damen et al. 2002, Popadic 2005), possibly extends to the crustacean limbs whenever uniramous, biramous or phyllopodous; reviewed in Williams (1999). In the ‘Uniramia’ concept, homology was assumed between the onychophoran jaw and the mandible of hexapods and myriapods, as opposed to that of the Crustacea (see below). It is now clear that the onychophoran jaw is not homologous with the mandible of myriapods, crustaceans and hexapods. The ‘Uniramia’ view has now generally been abandoned, see Shear (1992) for detailed discussion of morphological evidence. Most recent molecular phylogenies also support monophyly of the Arthropoda (e.g. Turbeville et al. 1991, Mallatt et al. 2004, Philippe et al. 2005, Dunn et al. 2008). Panarthropoda: Arthropods Plus Related Groups The clade grouping Tardigrada and Onychophora with Arthropoda has been called ‘Pan-Arthropoda’ (e.g. Nielsen 2001), meaning ‘all-arthropods’.

© 2011 by Taylor and Francis Group, LLC

58

Key Transitions in Animal Evolution

However, neither tardigrades nor onychophorans have articulated limbs. A synapomorphy uniting these three phyla is the presence of claws at the distal end of their limbs, as reflected in the word ‘Onychophora’ (i.e. ‘bearing nails’). Both Nielsen (2001) and Brusca and Brusca (2003) consider Tardigrada and Arthropoda as sister-groups on the basis of morphological synapomorphies, some of which, as articulated limbs with intrinsic muscles, are debatable (see Dewell et al. 1993, p. 144). Contrary to previous claims (Garey et al. 1996, Giribet et al. 1996), recent molecular phylogenies (Philippe et al. 2005, Baurain et al. 2007, Roeding et al. 2007, Timmermans et al. 2008) and a morphological study (Zantke et al. 2008) suggest that the Tardigrada could be closer to the Cycloneuralia (= Gastrotricha + Introverta). Similarity between the brains of onychophorans and cycloneuralians has also been proposed (Eriksson and Budd 2000). Several morphological characters, a heart with ostia and a pericardial septum, formation of the coelom, metanephridia, support a closer relationship between Onychophora and Euarthropoda than between the latter and Tardigrada (Schmidt-Rhaesa 2001). In a recent phylogenomic study (Dunn et al. 2008), Onychophora are closer to the Arthropoda with high support, while the position of Tardigrada, within Cycloneuralia or Pan-arthropoda depends on the phylogenetic method. On the other side, mitochondrial genes’ arrangements support the Tardigrada being closer to the Arthropoda than to the Nematoda (Ryu et al. 2007). At present, considering contradictory arguments, the relative position of Onychophora, Tardigrada and Arthropoda is not yet resolved. It has been suggested that Cycloneuralia are paraphyletic, that is, a monophyletic Pan-arthropoda would be issued from a cycloneuralian ancestor (Garey 2001, Mallatt et al. 2004). In my opinion, this hypothesis is most congruent with both molecular and morphological data. Noteworthy, Cycloneuralia paraphyly implies that the common ancestor of the Arthropoda and of the remaining cycloneuralians presented neither a ganglionic ventral nerve cord nor segmentation, characters that might be correlated (Schmidt-Rhaesa 2001). Are Pycnogonida the Sister-Group of the Euarthropoda? ‘Euarthropoda’, meaning ‘true’ arthropods, comprises Chelicerata, Myriapoda, Hexapoda and Crustacea plus all their fossils and their stem lineage fossils by opposition to some arthropod or arthropod-like fossils. They can be distinguished from the latter and from the Onychophora by loss of the appendage linked to the anterior-most part of the brain, the protocerebrum (Scholtz and Edgecombe 2006). The position of the Pycnogonida (= Pantopoda) (sea spiders) has been a matter of issue (see Dunlop and Arango 2005 for review). Indeed, Zrzavy et al (1997),

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

59

Fig. 2. Arthropod legs. A. Second leg of a grass hopper (Orthoptera, Hexapoda); B. A crustacean biramous limb (Anaspides, Malacostraca, Crustacea); C. Leg of Euryurus (Diplopoda, Myriapoda); D. Leg of a solpugid (Arachnida, Chelicerata); E. Leg of a sea spider (Pycnogonida, Chelicerata). (not to scale) Cx: coxa; Cxp: coxopodite; Eppds: epipodites; Expd: exopod; Fm; femur; Pat: patella; Ptar: pretarsus; Seg: body segment; Tar: tarsus; Tb: Tibia; Tr: trochanter; Tr1: first trochanter; Tr2: second trochanter. 1, 2, 3: proximal leg segments (in pycnogonid). All adapted from Snodgrass (1935).

© 2011 by Taylor and Francis Group, LLC

60

Key Transitions in Animal Evolution

Fig. 3. Homology of anterior segments in arthropods. Hexapoda: No appendage corresponding to the tritocerebral segment. Crustacea: In many species, the first legs of the thorax (or pereion) are transformed in a pair of maxillipedes, i.e. maxilla-like appendages. Chilopoda: No appendage corresponding to the tritocerebral segment; the first appendage of the trunk is transformed in a pair of forcipules (poison claws). Diplopoda: No appendage corresponding to the tritocerebral segment; the first and second pairs of maxillae are fused; no appendage in the first trunk segment. Arachnida: The deutocerebral segment bears a pair of chelicerae, the tritocerebral segment a pair of pedipalps; Arachnids have four pairs of walking legs. Pycnogonida: The first pair of legs is transformed in a pair of ovigers. Taking into account the ovigers as modified legs, pycnogonids have more legs than arachnids. a: antennae; a1: first pair of antennae; a2: second pair of antennae; ch: chelicerae; e: eyes; f: forcipules; l: labium; L: legs; mn: mandibles; mx: maxillae; mx1: first pair of maxillae; mx2: second pair of maxillae; o: ovigers; p: pedipalps. Color image of this figure appears in the color plate section at the end of the book.

Giribet et al. (2001) and Edgecombe (2004) placed them, although with weak support, as sister-group of all Euarthropoda on the basis of a combined molecular plus morphological phylogeny. This was supported by recent data from Maxmen et al. (2005) who claimed that the chelicerae (meaning ‘horns in the form of pincers’) of the pycnogonids belong to the protocerebral segment, thus excluding them from the Eu-arthropoda. Both morphological (discussed in Manuel et al. 2006) and molecular (Mallatt et

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

61

al. 2004) data contradict this view. In addition, expression of Hox genes (Manuel et al. 2006, Jager et al. 2006) clearly show that pycnogonids’ chelicerae are appendages belonging to the deutocerebral segment, homologous to those of ‘true’ chelicerates. Most systematists distinguish the Pycnogonida from all other chelicerates: the group (Pycnogonida + or Euchelicerata) is then called ‘Cheliceriformes’. In my opinion, the supposedly synapomorphies of the Euchelicerata, i.e. a prosomal carapace and genitals located in the first or second opisthosomal segment (Brusca and Brusca 2003, p. 697) can be interpreted as characters lost in pycnogonids in relation with their highly modified, and likely derived, body plan. Moreover, molecular phylogenies based on mitochondrial DNA retrieve Pycnogonida, not as a sister-group of all other chelicerates, i.e. Xiphosura (horseshoe crabs) and Arachnida, but instead nested within the Arachnida, as companions of Acari, Scorpiones, Aranae and others (Hassanin et al. 2005, Hassanin 2006, Park et al. 2007). Monophyly vs. Polyphyly of the Myriapoda ? Monophyly of the Myriapoda has been debated (Dohle 1997, Strausfeld 1998, Kraus 2001). The Myriapoda comprise four classes, two ‘major’ ones, i.e. Chilopoda (centipedes) and Diplopoda (millipedes), and two ‘minor’ ones, comprising less species, also of smaller size, the Symphyla and the Pauropoda. The latter two are usually united with the Diplopoda into the Progoneata clade. The heart of the issue is that no convincing

Fig. 4. Mandibulata vs. Paradoxopoda. A. Phylogeny of the Arthropoda under the Mandibulata hypothesis; B. Phylogeny of the Arthropoda under the Paradoxopoda hypothesis.

© 2011 by Taylor and Francis Group, LLC

62

Key Transitions in Animal Evolution

morphological synapomorphy unites the Chilopoda with the other three myriapod classes (Dohle 1997). In addition, cladistic analyses of brain characters resulted in ‘Myriapoda’ being a biphyletic assemblage, with Diplopoda appearing as the earliest branch of the arthropod tree and Chilopoda as the sister-group of the other mandibulates (Strausfeld 1998, Loesel et al. 2002). In contrast, the pattern of serotonergic neurons of the Diplopoda displays a closer similarity to that of the Chilopoda than to that of any other arthropod group (Harzsch 2004, p. 209). From an extensive cladistic analysis of morphological and molecular characters, Edgecombe (2004) concluded in favour of the monophyly of Myriapoda, and underlined several synapomorphies, such as the structure of the tentorium (a piece of the cephalic skeleton), the secondary division of the gnathobasic mandible in two pieces, with corresponding muscles. On the molecular side, most recent phylogenies with various molecules and methods consistently retrieve a myriapod clade comprising both millipedes and centipedes (Elongation factor 2: Regier and Shultz 2001; both 18S and 28S rRNA: Mallatt et al. 2004; EF1-alpha: Pisani 2004; 9 nuclear and 15 mitochondrial genes: Pisani et al. 2004; three nuclear protein genes: Regier et al. 2005; mitochondrial DNA: Hassanin 2006, contra Negrisolo et al. 2004); see discussion of both morphological and molecular phylogenies in Bitsch and Bitsch 2004). Mandibles: Paradoxopoda vs. Mandibulata As shown by their name, chelicerates bear chelicerae related to the deutocerebrum, whereas the other three sub-phyla bear a pair of antennae at the same location. In addition to a pair of antennae on the deutocerebral segment, all three groups share a pair of mandibles on the first postoral (gnathal) segment, whereas the chelicerates exhibit their first pair of legs, hence the name ‘Mandibulata’ for the grouping of Myriapoda, Crustacea and Hexapoda. Homology of the anterior segments between the Chelicerata and Mandibulata is now conclusively supported by evidence for the presence of a deutocerebrum in chelicerates (Mittmann and Scholtz 2003), contrary to previous thoughts (e.g. Bullock and Horridge 1965, vol. II, chap. 16, p. 821, Wegerhoff and Breidbach 1995), and from the expression of Hox genes in chelicerates belonging to various groups (Telford and Thomas 1998, Damen et al. 1998, Jager et al. 2006). In addition, it is now clear that all cephalic appendages are serially homologous to the legs. This is based in particular on the use of a common network of developmental genes to build these appendages (Casares and Mann 1998, 2001, Ronco et al. 2008). Hence, a primitive leg-like appendage corresponding to the deuterocerebral segment (Mittmann and Scholtz 2003, Scholtz and Edgecombe 2006) has evolved as a chelicera in chelicerates (Abzhanov

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

63

and Kaufman 2000, Prpic and Damen 2004) and as an antenna in the other arthropod sub-phyla. In addition, a primitive leg-like appendage has been transformed into a mandible altogether in myriapods, crustaceans and hexapods. Homology between mandibles of these three groups is not only supported by their segmental position, but also by their detailed structure (Edgecombe et al. 2003), which makes convergent evolution unlikely. However, the clade ‘Mandibulata’ has recently been rejected by molecular phylogenies based on various molecules: hemocyanin (Kusche and Burmester 2001); 28S and 18S ribosomal RNA (Mallatt et al. 2004); mitochondrial DNA (Hwang et al. 2001, Hassanin et al. 2005, Hassanin 2006), Hox genes (Cook et al. 2001), various nuclear genes (Pisani et al. 2004). Indeed, Myriapoda, instead of grouping with Crustacea and Hexapoda, were found as sister-group of the Chelicerata. This clade was coined ‘Paradoxopoda’ by Mallatt et al. (2004), also called ‘Myriochelata’ (Pisani et al. 2004). To my knowledge, only a few developmental-morphological similarities have been brought in support to the Paradoxopoda. In both Myriapoda and Chelicerata neurogenesis does not proceed with the formation of single neuroblasts as in Hexapoda and Crustacea (Gerberding 1997, Harzsch et al. 1998, Gerberding and Scholtz 2001, Harzsch 2001, Ungerer and Scholtz 2008), but by clusters of neurogenic cells, (Mittmann 2002, Stollewerk 2002, Stollewerk et al. 2003, Dove and Stollewerk 2003, Kadner and Stollewerk 2004, Stollewerk and Simpson 2005, Stollewerk and Chipman 2006). The structure of midline brain neuropils is similar in chelicerates and chilopods (but not diplopods) (Strausfeld 1998, Loesel et al. 2002). However, these traits may well be plesiomorphic, not shared derived properties, thus with no value to unite Myriapoda and Chelicerata in a convincing clade (Strausfeld 1998, Harzsch et al. 2005, Stollewerk and Chipman 2006, Stollewerk 2008). Following the Paradoxopoda hypothesis, the antenna, the mandible and the maxillae found in both Myriapoda and Tetraconata (= Crustacea + Hexapoda) have to be interpreted not as homologous organs, but due to evolutionary convergences. Prpic and Tautz (2003) argued that the gnathobasic mandible could be convergent between myriapods and tetraconates in view that the maxilla also is gnathobasic in myriapods but is not in crustaceans and hexapods. However, this leaves aside the fact that in chelicerates, the corresponding appendages are neither gnathobasic nor telobasic mouthparts, they are legs actually. Hence, the myriapod mandible and maxilla are both derived appendages, and the tetraconate mandible and maxilla are derived as well. This does not exclude convergence, but does not allow comparison between the fate of mandibles and that of maxillae. Another character of interest is provided by the leg’s architecture. Although all arthropod legs are based on a roughly similar plan (see above), an additional podomere, the ‘patella’, is intercalated between

© 2011 by Taylor and Francis Group, LLC

64

Key Transitions in Animal Evolution

femur and tibia in chelicerates. Pycnogonid legs also possess a patella, but differ from the other chelicerates’ legs by additional segments at the proximal end (Snodgrass 1935, Chap. 5). In all other arthropods, a patella is absent from legs. Phylogenetic interpretation of this character is equivocal. If the arthropod primitive state does not include a patella, it can be viewed as a synapomorphy of the Chelicerata (including Pycnogonida). If the presence of a patella is plesiomorphic, its loss can be taken either as a synapomorphy of the Mandibulata or as a homoplasy. We are thus faced to severe discrepancies between molecules and morphology. As there is but a single evolutionary history, one of the two evolutionary hypotheses, ‘Mandibulata’ vs. ‘Paradoxopoda’, must be wrong. In recent papers, Telford et al. (2008) and Rota-Stabelli and Telford (2008) showed that the molecular phylogeny of the diverse phyla and classes among the Panarthropoda strongly depends on the outgroup choice. When the Priapulida, a member of the Cycloneuralia with slow evolutionary rate, is taken as outgroup, then molecular phylogenies support the Mandibulata, reconciling molecular with morphological evidence. Hexapods are Terrestrial Crustaceans The idea that the Crustacea could be paraphyletic can be traced back as early as 1904 (Lankester 1904). E. Ray Lankester was the first to classify horseshoe crabs (Limulus) not within the Crustacea, as previously thought, but within the Chelicerata. In his view, all mandibulates were issued from a crustacean ancestor. More recently, Briggs and Fortey (1989), on the basis of a cladistic analysis of Cambrian and extant arthropods, pushed paraphyly in as much as to group all arthropod classes, including the Trilobita and the Chelicerata, as issued from a crustacean-like ancestor. However, this view received little agreement among zoologists, until the paraphyly of the Crustacea was resurrected in its present avatar: inclusion of the Hexapoda only within the Crustacea. Reviews and debates can be found in Fortey and Thomas (1997) and Deuve (2001). The so-called ‘Atelocerata’ or ‘Tracheata’ assemblage (Myriapoda + Hexapoda) was supposed to be supported by morphological traits, such as tracheae and Malphighian tubules, absent from the Crustacea. However, these characters are thought to be convergences due to terrestrial life. Indeed, similar structures are also found in other panarthropod groups. Loss of the second antenna could be homoplastic, as often for character loss. The last character of contention was the mandible, which would be ‘gnathobasic’ in crustaceans, biting with the base of the mandibles, whereas myriapods and insects would have whole-limb mandibles that bite with their tips (so-called ‘telognathic’) (Fryer 1996 p. 2–7). However,

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

65

recent works, using an antibody against the Distal-less protein known to be involved in appendage development (Panganiban et al. 1997), on a variety of myriapod, crustacean and hexapod species (Popadic et al. 1996, 1998, Scholtz et al. 1998, Prpic 2004), showed that mandibles of all mandibulates are of the gnathobasic type, thus supporting their homology (Scholtz 2001). This led to the demise of the ‘Atelocerata’ (Telford and Thomas 1995; but see in contrast Bitsch and Bitsch 2004). At the same time, molecular phylogenies began to give support to a closer relation between Crustacea and Hexapoda than between the latter and Myriapoda (Turbeville et al. 1991, Boore et al. 1995, Friedrich and Tautz 1995). These first studies were based on a small amount of data and a very small sample of arthropod species. The close relation between hexapods and crustaceans has later been vindicated by numerous studies on a variety of taxa and molecules: ribosomal RNAs (Mallatt et al. 2004); various nuclear coding genes (Shultz and Regier 2000, Regier and Shultz 2001, Cook et al. 2001, Giribet et al. 2001, Pisani 2004, Baurain et al. 2007); mitochondrial DNA (Boore et al. 1998, Garcia-Machado et al., 1996, 1999, Wilson et al. 2000, Hwang et al. 2001, Lavrov et al. 2004, Cook et al. 2005, Hassanin 2006, Lim and Hwang 2006). A number of morphological/developmental characters are shared by hexapods and crustaceans at the exclusion of chelicerates and myriapods. Most of them are related to the nervous system. In both Hexapoda and Crustacea, neurogenesis proceeds with the primary formation of neuroblasts, which are stem cells giving rise to neurons, glia and sensory cells. Moreover, based on their origin and position, some individual neuroblasts (Ungerer and Scholtz 2008) and a number of pioneer neurons are clearly homologous in both groups (Whitington 1995, 1996). They give rise to homologous nerve tracts (Harzsch 2004). At least some ganglia and neuropils composing the brain are also homologous (in addition to papers published in Fortey and Thomas (1997) and Deuve (2001), see Osorio and Bacon 1994, Osorio et al. 1995, Strausfeld 1998). In this respect, the visual system (eyes and optic lobes) is of particular interest. Looking at Arthropod Eyes: Homology of Arthropod Compound Eyes ? Opticians distinguish ‘conventional’ cameras with a single aperture to ‘compound’ ones, with multiple apertures. The facetted compound eye of arthropods seems such an extraordinary organ as it deserves particular attention. Compound eyes are found in all four sub-phyla of the Arthropoda. However, they are found in a single group of chelicerates, the Xiphosura (horse-shoe crabs, e.g. Limulus), and in a single group of myriapods, the Scutigeromorpha (a family of chilopods, e.g. the centipede Scutigera). On the opposite, leaving aside blind animals, almost all crustaceans and hexapods have compound eyes.

© 2011 by Taylor and Francis Group, LLC

66

Key Transitions in Animal Evolution

Reciprocally, not all arthropod eyes are compound eyes. In addition to compound facetted eyes, simple eyes are present in arthropods. ‘Simple’ does not mean an eye with a single photoreceptor unit. Our own eyes are simple eyes; still our retina contains millions of photoreceptor cells. The simple ventral eye of the horseshoe crab Limulus contains about 300 photoreceptor cells (Battelle 2006). The ‘simple’ frontal eyes of wolf spiders possess quite a sophisticated optic apparatus (Williams and McIntyre 1980). Besides simple and compound eyes, I distinguish a third type that I call ‘multiple eyes’. They are formed by groups of ocelli (small eyes). They are diversely located, at the lateral sides of the head as the stemmata of holometabolous insect larvae, or at the dorsal side as the median eye of the crustacean nauplius larva. These three different types of eyes may be found in the same species, either simultaneously or at different stages of its life cycle. It is far beyond the scope of the present chapter to review in detail the diversity of arthropod eyes. In the following sections, I will focus on a single issue, related to the present topic: Are compound eyes homologous throughout the Arthropoda? The classical view posits that the compound eye is a synapomorphy of the Arthropoda (e.g. Nielsen 2001). By itself the diversity of arthropod eyes and their dispatching between the various taxonomic groups question this hypothesis: homology implies that the common ancestor of all arthropods possessed a compound eye, then, loss would have happened a number of times during the evolution of the Arthropoda. Conversely, non-homology implies that compound eyes would have been designed de novo several times. The basic unit of compound eyes is the ommatidium, a complex structure comprising several photoreceptor neurons (retinula cells) and other components such as non-photoreceptor pigment cells, lenses and accessory cells. Although by etymology ‘ommatidium’ means ‘small eye’, a compound eye is not the mere juxtaposition of small simple eyes. The intricate complexity of neuronal pathways of in the optic lobes as studied in Drosophila (Meinertzhagen and Hanson 1993) shows that vision is the result of integration of inputs from several ommatidia, if not from the whole eye. In other words, an ommatidium isolated from a compound eye cannot ‘see’ anything by itself. To discuss homology between the various arthropod compound eyes, it is thus necessary to examine the relations between the different elements of the ommatidium and the neuronal connections between retina and brain.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

Drosophila cornea cone

Scutigera cornea pigment cell

cone cell

cone

pigment cell rhabdome

distal retinula cell distal rhabdome

R7 rhabdomere retinula cell

proximal rhabdome proximal retinula cell

67

Limulus cornea

pigment cell

rhabdome

retinula cell R8 rhabdomere

eccentric cell

axons

Fig. 5. Ommatidia. A. The tetraconate ommatidium: ommatidium of the Drosophila eye; simplified from Wolff and Ready (1993); B. Ommatidium of the Scutigera eye; adapted from Paulus (2000); C. Ommatidium of the Limulus eye, adapted from Nilsson and Kelber (2007). (not to scale).

The structure of the ommatidium in compound eyes Ommatidia from crustacean and hexapod compound eyes are strikingly similar (Melzer et al. 1997, Dohle 2001). Each ommatidium contains eight retinula cells (neuronal photoreceptor cells), in which the accumulation of opsin forms a rhabdome along the proximal to distal axis. At the distal part of the ommatidium, under the cornea, a crystalline cone forms a lens, produced by four cone cells. This led Dohle to propose the name ‘Tetraconata’ for the clade [Crustacea + Hexapoda] (Dohle 2001, Richter 2002), also called ‘Pancrustacea’. This pattern of four cone cells and eight retinula cells is fairly constant in hexapods and malacostracans, but some minor variations can be found in non-malacostracan crustaceans (Hallberg and Elofsson 1983, Dohle 2001, Richter 2002, Oakley and Cunningham 2002, reviewed in Bitsch and Bitsch 2005, discussion in Nilsson and Kelber 2007). Paulus (2000, p. 190) concluded: “the ommatidia of the Crustacea and Insecta are clearly homologous”. Structural similarities of the crustacean and hexapod eyes result from closely similar developmental processes (reviewed in Harzsch and Hafner 2006). The parallel between crustacean

© 2011 by Taylor and Francis Group, LLC

68

Key Transitions in Animal Evolution

and insect eye formation has been extended to the developmental genetic level by the similar role of the wingless gene in eye growth in Drosophila and in the malacostracan crustacean Mysidium (Duman-Scheel et al. 2002). The structure of the compound eye of scutigeromorph myriapods has been reviewed and revisited in detail by Müller et al (2003). The number and composition of cell types varies within the eye (central to marginal). Like the crustacean and hexapod ommatidia, Scutigera’s ommatidium comprises a crystalline cone made by four (sometimes five) cone cells. On this respect it deserves the name of ‘tetraconate’. It differs from the pancrustacean ommatidium by a two-layered rhabdome: a distal open rhabdome made by 9 to 12 retinulae and a proximal closed rhabdome made by four retinula cells, instead of the single-layered rhabdome made by 8 retinula cells in Pancrustacea. Inter-ommmatidial cells are present. Some of which are pigment cells that isolate ommatidia from each other as in crustacean/hexapod eyes, while others are glandular cells, specific to scutigeromorph eyes. Summarising, the ommatidium of the scutigeromorph compound eye, although sharing some similarities, presents peculiar features that largely outcome the range of variations seen among ommatidia of crustaceans and hexapods. The structure of the xiphosuran ommatidium is quite different (Fahrenbach 1968, 1969, reviewed in Paulus 2000, Bitsch and Bitsch 2005, Battelle 2006). First, there is no typical crystalline cone. Indeed, there is no differentiation between the cornea and the ‘cone’. The cone of the Limulus’ ommatidium is but an expansion of the cuticular cornea (Fahrenbach 1968). The number of retinula cells is variable, from 4 to as many as 20 (average 10–13) per ommatidium (Fahrenbach 1969). These retinula cells form a rhabdome in the centre of which a neuron, so-called ‘eccentric cell’ is located (sometimes two of them when the number of rhabdomeric cells is high). This eccentric cell is never seen in other arthropod compound eyes. The arrangement of the ommatidia is variable, not forming a regular hexagonal pattern on the whole surface of the eye (Fahrenbach 1969, Harzsch et al. 2006). These structural aspects result from differences in development (Harzsch et al. 2006, Harzsch and Hafner 2006). Summarising, the ommatidium of the compound eye of the Xiphosura is so different from those of the other arthropods that, on both structural and developmental criteria, it is likely evolutionary convergent. This is in agreement with both the ‘Paradoxopoda’ and the ‘Mandibulata’ phylogenetic hypotheses. The great similarity of the ommatidium between Crustacea and Hexapoda brings support to the ‘Tetraconata/Pancrustacea’ hypothesis. The scutigeromorph ommatidium presents some similarities with the pan-crustacean one. This leaves the position of the Myriapoda equivocal: if the scutigeromorph compound eye and ommatidium are thought to be part of the myriapod ground plan, this would favour the

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

69

‘Mandibulata’ (with loss of the compound eye in the other myriapod groups), whilst if on the opposite they are taken as scutigeromorph apomorphies within the Myriapoda, these characters will be of no value in the ‘Paradoxopoda’ vs. ‘Mandibulata’ issue. Optic Lobes in Crustaceans and Hexapods Hanström (1947, p. 42) underlined the similarity between the pattern of neuropils and nerve nets in hexapod and malacostracan optic lobes and in more recent times Osorio drew attention to its importance for phylogenetic relationships (Osorio and Bacon 1994, Osorio et al. 1995, Nilsson and Osorio 1997). Both groups of arthropods possess (at least) three neuropils that directly or indirectly receive neuronal inputs from the retina, i.e. going from the distal side (retina) to the proximal one (protocerebrum proper): lamina, medulla and lobula. Nerve tracks between these neuropils form two chiasmata, one between the lamina and the medulla and a second between the medulla and the lobula. Such an organization is never found in any other arthropod, including non-malacostracan crustaceans, socalled ‘entomostracans’. Indeed, the latter have only two neuropils with no neuronal crossing in between.

Fig. 6. Optic lobes: neuropils and projections from the retina. Scutigeromorpha: a family of chilopods (centipedes), see text; Branchiopoda: a class of nonmalacostracan crustaceans, e.g. Artemia, Triops, Daphnia; Decapoda: the most numerous class of malacostracan crustaceans, e.g. lobsters, crabs, crayfish; Pterygota: winged insects. In Scutigeromorphs and in non-malacostracan crustaceans, the lamina, the first optic neuropil, is followed by an unique second neuropil, with no chiasma in between. In malacostracans and in hexapods, the lamina is followed through a chiasma by the medulla (divided in medulla interna and medulla externa in insects), which in turn is followed by the lobula through a second chiasma and by a fourth neuropil, the deep lobula (malacostracans) or lobula plate (insects). Simplified from Sinakevitch et al. (2003).

© 2011 by Taylor and Francis Group, LLC

70

Key Transitions in Animal Evolution

Sinakevitch et al. (2003) re-examined in detail the morphology of the optic lobes in a variety of crustaceans and insects. They confirmed that the architecture of the optic lobes of branchiopod crustaceans and of scutigeromorphs greatly differs from that of malacostracans and hexapods. The main outcome of this study is that, in addition to the ‘classical’ lamina, medulla and lobula, a fourth neuropil is retrieved in these latter two groups, called the lobula plate in dipteran insects and lobula satellite, sublobula or deep lobula in the other species. Despite some diversity, a common and likely homologous plan can be derived. Strikingly similar morphogenetic movements during development of the optic lobes in the malacostracan shrimp Penaeus (Elofsson 1969) and in the fly Drosophila (Meinertzhagen and Hanson 1993) result in the formation of chiasmata. The process of optic lobe development is different in branchiopods (Elofsson and Dahl 1970, Harzsch and Walossek 2001). Although convergence cannot be formally excluded (Bitsch and Bitsch 2005), their development and the relations between their various elements strongly argue in favour of homology of optic lobes between malacostracans and hexapods; for a more complete survey see Harzsch (2002). On the molecular phylogenetic side, a closer relationship between the Hexapoda and the Malacostraca is found only with mitochondrial DNA (Garcia-Machado et al. 1996, 1999, Wilson et al. 2000, Lim and Hwang 2006), not with phylogenies based on other molecules. Are Hexapoda Polyphyletic ? The Collembolan Controversy In 2003, Nardi et al. claimed that the ‘Hexapoda’ were a polyphyletic assemblage. Indeed, in their phylogenetic analysis of mitochondrial DNA, the Collembola (springtails) stemmed from the base of the Tetraconata clade, leaving the Crustacea and the rest of the hexapods as sister-groups (Fig. 7A). The collembolans, with proturans and diplurans, belonging to two other orders of wingless hexapods, have their mouth pieces located inside a cephalic cavity, contrary to all other hexapods, in which the mouth pieces can be seen outside the head capsule, thus forming the Ectognatha (= Insecta sensu stricto) clade. However, the relative positions of the three endognathous orders at the base of the hexapod phylogenetic tree are still unresolved or controversial. Nardi et al.’s analysis soon raised critiques about biases, mainly because of heterogeneity in base composition of the mtDNA sequences used (Delsuc et al. 2003). Later, Cook et al. (2005) phylogenetic analyses of mtDNA sequences also suggested a polyphyletic ‘Hexapoda’ but with a pattern different from that of Nardi et al.: Collembola branched this time not as a sister-group of all other tetraconates, but as a sister group to the

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

71

Fig. 7. The position of the Collembola within the Arthropoda. A. Polyphyletic hypothesis 1, based on mtDNA (simplified from Nardi et al. 2003, Fig. 2); B. Polyphyletic hypothesis 2, based on mtDNA (simplified from Cook et al. 2005, Fig. 2); C. Monophyletic hypothesis, based on ribosomal proteins (simplified from Timmermans et al. 2008, Fig. 2).

so-called ‘maxillopod’ crustaceans, while the ‘true’ Insecta grouped with branchiopod and malacostracan crustaceans (Fig. 7B). Data from mtDNA alone cannot resolve the relationships between Crustacea, Collembola and Insecta (Cameron et al. 2004), even when the compositional bias of the mtDNA is taken into account (Holland et al. 2005, Hassanin et al. 2005, Hassanin 2006). In contrast, ribosomal RNA-based phylogenies (Kjer 2004, Mallatt et al. 2004, Luan et al. 2005, Mallatt and Giribet 2006, Misof et al. 2007) and nuclear protein-coding genes (Regier et al. 2005, Timmermans et al. 2008) retrieve Hexapoda, including Collembola, monophyletic with high support (Fig. 7C). Development and Body Plan of the Collembolans In fact, the idea that collembolans do not belong to the Hexapoda was not new (see Kristensen 1995 and Bitsch et al. 2004 for references and discussion, with morphological data supporting hexapod monophyly). Whatsoever, the development and the body plan of the Collembola are worth examining. They are unique in presenting a total cleavage (Jura 1972), while the early development of all other hexapod species begins with superficial (partial) cleavage, the first divisions of the zygote forming a syncytial embryo. However, superficial cleavage is found in several crustacean species (Anderson 1973, p. 278), making the type of cleavage a homoplastic character (see discussion in Scholtz 1997). Collembolan embryos possess appendage buds not only on the thorax, but also on all abdominal segments. However, this again is not specific to collembolans,

© 2011 by Taylor and Francis Group, LLC

72

Key Transitions in Animal Evolution

the same being found in other hexapod orders, including within the Ectognatha, such as the Archeognatha (Jura 1972). Regarding their body plan, collembolans possess three pairs of legs, thus deserving the name of ‘hexapods’. This is not as self-evident as it may seem: indeed, although the crustacean classes present a wide panel of tagmata, segments with and/or without trunk appendages in varying numbers, there is no crustacean with three pairs of legs. More importantly, collembolans are unique among the Hexapoda in having at most six abdominal segments, whereas the basic hexapod number is 11 (Brusca and Brusca 2003). As adults, they bear a curious appendage, the collophore, on the first abdominal segment, another, the retinaculum, on the third abdominal segment, and on the fourth a special appendage, the furcula, used for jumping, hence their name of ‘springtails’. Summary and Conclusions In the present chapter, I have addressed the following issues: 1. What is the position of the Arthropoda in the animal kingdom? 2. Are Arthropoda monophyletic? 3. Which, between Tardigrada and Onychophora, is the sister group of the Euarthropoda? 4. Are Pycnogonida the sister-group of the Euarthropoda? 5. What are the relations between the four arthropod subphyla, i.e. ‘Paradoxopoda’ vs. ‘Mandibulata’? 6. Are Myriapoda monophyletic? 7. Are hexapods terrestrial crustaceans? 8. Are Hexapoda monophyletic? In my opinion, some of these questions have received an answer thanks to advances during the recent decades in phylogenetic analyses, in morphology due to new and improved techniques, and in comparative studies of development, including in their genetic aspects (‘evo-devo’). I favour those answers where there is congruence between molecules and morphology, ‘evo-devo’ being integration of both molecular and morphological studies. So, I think there is enough consistent data favouring the following answers: 1. Arthropoda is a member of the Ecdysozoa, probably issued from a cycloneuralian stem. 2. Arthropoda is a monophylum, with little doubt. 4. Pycnogonida is a member of the Chelicerata, not the sister-group of all other ‘Euarthropoda’. 5. Mandibulata is a monophyletic group, Myriochelata/Paradoxopoda is likely a methodological artefact in molecular analyses.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

73

7. Hexapoda is issued from a crustacean stem, thus making the ‘Crustacea’ sensu stricto a paraphyletic assemblage. 8. Hexapoda is a monophylum, claims for polyphyly are likely due to reconstruction artefacts in molecular phylogenies. Some questions will hopefully receive a clear answer within the next few years: 3. Which, Onychophora or Tardigrada, is the sister group of Euarthropoda? At present, this issue is still open. Answering requires morphological investigations on more species, molecular analyses with more species and molecules using as out-groups members of the Cycloneuralia. 4. Are Pycnogonida the sister-group of all other chelicerates or are they nested within the Arachnida? The precise position of the Pycnogonida needs to be re-examined on a molecular ground using various molecules, not only mitochondrial, more sea-spider species, more other chelicerate species and appropriate out-groups. Morphological characters have also to be evaluated more precisely with more data using modern methods. Whatsoever, there is little doubt that Pycnogonida do belong to the Chelicerata, and thus the term ‘Cheliceriformes’ is obsolete. 6. Are the Myriapoda monophyletic? In my opinion, the question is still open. More studies are needed, with morphological, molecular and evo-devo approaches, on more ‘myriapod’ species, combined with parallel studies on closer and multiple outgroups. 7. Are Malacostraca the closest relatives to the Hexapoda? Morphology and development, most of the nervous and ocular systems, provide convincing data for such a relation. However, it is congruent only with mitochondrial DNA-based molecular phylogenies, which is unsatisfactory. It should be said that molecular phylogenies have been up to now desperately unable to resolve with confidence, because of lack of resolution and/or inconsistency between the various analyses/authors, the relationships not only between hexapods and crustaceans, but also between the various crustacean groups. Endeavour in this field is awaited. Acknowledgements Many thanks to Gerhard Scholtz for his comments and advice on a previous draft of this manuscript and to Louis Deharveng and Cyrille D’Haese for reading and help on the collembolan section.

© 2011 by Taylor and Francis Group, LLC

74

Key Transitions in Animal Evolution

References Abu-Shaar, M. and R.S. Mann. 1998. Generation of multiple antagonistic domains along the proximodistal axis during Drosophila leg development. Development 125: 3821–3830. Abzhanov, A. and T.C. Kaufman. 2000. Homologs of Drosophila appendage genes in the patterning of arthropod limbs. Dev. Biol. 227: 673–689. Adoutte, A. and G. Balavoine, N. Lartillot, O. Lespinet, B. Prud’homme, and R. de Rosa. 2000. The new animal phylogeny: reliability and implications. Proc. Natl. Acad. Sci. USA 97: 4453–4456. Aguinaldo, A.M. and J.M. Turbeville, L.S. Linford, M.C. Rivera, J.R. Garey, R.A. Raff, and J.A. Lake. 1997. Evidence for a clade of nematodes, arthropods and other moulting animals. Nature 387: 489–493. Amundson, R. Richard Owen and animal form. pp. XV-LI. In: R. Amundson. [ed.] 2007. R. Owen. On the nature of limbs. Chicago University Press, Chicago. Anderson, D.T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon Press, Oxford. Averof, M. and S.M. Cohen. 1997. Evolutionary origin of insect wings from ancestral gills. Nature 385: 627–630. Ballard, J.W.O. and G.J. Olsen, D.P. Faith, W.A. Odgers, D.M. Rowell, and P.W. Atkinson. 1992. Evidence from 12S ribosomal RNA sequences that Onychophorans are modified arthropods. Science 258: 1345–1348. Battelle, B.A. 2006. The eyes of Limulus polyphemus (Xiphosura, Chelicerata) and their afferent and efferent projections. Arthropod. Struct. Dev. 35: 261–274. Baurain, D. and H. Brinkmann, and H. Philippe. 2007. Lack of resolution in the animal phylogeny: closely spaced cladogeneses or undetected systematic errors? Mol. Biol. Evol. 24: 6–9. Bitsch, C. and J. Bitsch. 2004. Phylogenetic relationships of basal hexapods among the mandibulate arthropods: a cladisitic analysis based on comparative morphological characters. Zool. Script. 33: 511–550. Bitsch, C. and J. Bitsch. Evolution of eye structure and arthropod phylogeny. pp. 185–214. In: S. Koenemann and R.A. Jenner. [eds.] 2005. Crustacea and Arthropod Relationships. CRC Press, Boca Raton, USA. Boore, J.L. and T.M. Collins, D. Stanton, L.L. Daehler, and W.M. Brown. 1995. Deducing the pattern of arthropod phylogeny from mitochondrial DNA rearrangements. Nature 376: 163–165. Boore, J.L. and D.V. Lavrov, and W.M. Brown. 1998. Gene translocation links insects and crustaceans. Nature 392: 667–668. Briggs, D.E.G. and R.A. Fortey. 1989. The early radiation and relationships of the major arthropod groups. Science 246: 241–243. Brusca, R.C. and G.J. Brusca. 2003. Invertebrates. Sinauer, Sunderland MA, USA. Bullock, T.H. and G.A. Horridge. 1965. Structure and Function in the Nervous System of Invertebrates. Freeman & Co., San Francisco, USA. Casares, F. and R.S. Mann. 1998. Control of antennal versus leg development in Drosophila. Nature 392: 723–726. Casares, F. and R.S. Mann. 2001. The ground state of the ventral appendage in Drosophila. Science 293: 1477–1480. Cook, C.E. and M.L. Smith, M.J. Telford, A. Bastianello, and M. Akam. 2001. Hox genes and the phylogeny of the arthropods. Curr. Biol. 11: 759–763. Cook, C.E. and Q. Yue, and M. Akam. 2005. Mitochondrial genomes suggest that hexapods and crustaceans are mutually paraphyletic. Proc. Biol. Sci. 272: 1295–1304. Costello, D.P. and C. Henley. 1976. Spiralian development: A perspective. Am. Zool. 16: 277–291. Cuvier, G. 1817. Le règne animal, distribué d’après son organisation, pour servir de base à l’histoire naturelle et d’introduction à l’anatomie comparée. Deterville, Paris, France.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

75

Damen, W.G. and M. Hausdorf, E.A. Seyfarth, and D. Tautz. 1998. A conserved mode of head segmentation in arthropods revealed by the expression pattern of Hox genes in a spider. Proc. Natl. Acad. Sci. USA 95: 10665–10670. Damen, W.G. and T. Saridaki, and M. Averof. 2002. Diverse adaptations of an ancestral gill: a common evolutionary origin for wings, breathing organs, and spinnerets. Curr. Biol. 12: 1711–1716. Darwin, C. 1872. The Origin of Species. Murray, London, UK. de Beer, G.R. Embryology and evolution. pp. 57–78. In: G.R. de Beer. [ed.] 1938. Evolution. Essays on Aspects of Evolutionary Biology. Clarendon Press, Oxford, UK. de Rosa, R. and J. Grenier, T. Andreeva, C. Cook, A. Adoutte, M. Akam, S. Carroll, and G. Balavoine. 1999. Hox genes in brachiopods and priapulids and protostome evolution. Nature 399: 772–776. Deuve, T. 2001. The origin of the Hexapoda. Ann. Soc. Entomol. Fr. vol 37. Paris, France. Dewell, R.A. and D.R. Nelson, and W.C. Dewell. Tardigrada. pp. 143–183. In: F.W. Harrison [ed.] 1993. Microscopic Anatomy of Invertebrates. Wiley-Liss, New York. Dohle, W. Myriapod-insect relationships as opposed to an insect-crustacean sister group relationship. pp. 305–315. In: R.A. Fortey and R.H. Thomas. [eds.] 1997. Arthropod Relationships. Chapman & Hall, London. Dohle, W. 2001. Are the insects terrestrial crustaceans? A discussion of some new facts and arguments and the proposal of the proper name ‘Tetraconata’ for the monophyletic unit Crustacea + Hexapoda. Ann. Soc. Entomol. Fr. 37: 85–103. Dove, H. and A. Stollewerk. 2003. Comparative analysis of neurogenesis in the myriapod Glomeris marginata (Diplopoda) suggests more similarities to chelicerates than to insects. Development 130: 2161–2171. Duman-Scheel, M. and N. Pirkl, and N.H. Patel. 2002. Analysis of the expression pattern of Mysidium columbiae wingless provides evidence for conserved mesodermal and retinal patterning processes among insects and crustaceans. Dev. Genes Evol. 212: 114–123. Dunlop, J.A. and C.P. Arango. 2005. Pycnogonid affinities: a review. J. Zool. Syst. Evol. Research 43: 8–21. Dunn, C.W. and A. Hejnol, D.Q Matus, K. Pang, W.E. Browne, S.A. Smith, E. Seaver, G.W. Rouse, M. Obst, G.D. Edgecombe, et al. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–749. Edgecombe, G.D. and S. Richter, and G.D.F. Wilson. 2003. The mandibular gnathal edges: Homologous structures throughout Mandibulata? African Invert. 44: 115–135. Edgecombe, G.D. 2004. Morphological data, extant Myriapoda and the myriapod stem group. Contrib. Zool. 73 (3): 2. Elofsson, R. 1969. The development of the compound eyes of Penaeus duorarum (Crustacea: Decapoda) with remarks on the nervous system. Z. Zellforsch. Mikrosk. Anat. 97: 323–350. Elofsson, R. and E. Dahl. 1970. The optic neuropiles and chiasmata of Crustacea. Z. Zellforsch. Mikrosk. Anat. 107: 343–360. Elofsson, R. 2006. The frontal eyes of crustaceans. Arthropod. Struct. Dev. 35: 275–291. Eriksson, B.J. and G.E. Budd. 2000. Onychophoran cephalic nerves and their bearing on our understanding of head segmentation and stem-group evolution of Arthropoda. Arthropod. Struct. Dev. 29: 197–209. Fahrenbach, W.H. 1968. The morphology of the eyes of Limulus. I. Cornea and epidermis of the compound eye. Z. Zellforsch. 87: 278–291. Fahrenbach, W.H. 1969. The morphology of the eyes of Limulus. II. Ommatidia of the compound eye. Z. Zellforsch. 93: 451–483. Fortey, R.A. and R.H. Thomas. 1997. Arthropod Relationships. The Systematics Association. vol. 55. Chapman & Hall, London, UK. Friedrich, M. and D. Tautz. 1995. Ribosomal DNA phylogeny of the major extant arthropod classes and the evolution of myriapods. Nature 376: 165–167. Friedrich, M. 2006. Continuity versus split and reconstitution: exploring the molecular developmental corollaries of insect eye primordium evolution. Dev. Biol. 299: 310–329.

© 2011 by Taylor and Francis Group, LLC

76

Key Transitions in Animal Evolution

Fryer, G. 1996. Reflections on arthropod evolution. Biol. J. Linnean Soc. 58: 1–55. Fryer, G. A defence of arthropod polyphyly. pp. 23–33. In: R.A. Fortey and R.H. Thomas. [eds.] 1997. Arthropod Relationships. Chapman & Hall, London, UK. Garcia-Machado, E. and N. Dennebouy, M.O. Suarez, J.C. Mounolou, and M. Monnerot. 1996. Partial sequence of the shrimp Penaeus notialis mitochondrial genome. C. R. Acad. Sci. Paris 319: 473–486. Garcia-Machado, E. and M. Pempera, N. Dennebouy, M. Oliva-Suarez, J.C. Mounolou, and M. Monnerot. 1999. Mitochondrial genes collectively suggest the paraphyly of Crustacea with respect to Insecta. J. Mol. Evol. 49: 142–149. Garey, J.R. and M. Krotec, D.R. Nelson, and J. Brooks. 1996. Molecular analysis support a tardigrade-arthropod association. Invert. Biol. 115: 79–88. Garey, J.R. 2001. Ecdysozoa: The relationship between Cycloneuralia and Panarthropoda. Zool. Anz. 240: 321–330. Geoffroy Saint-Hilaire, E. 1818. Philosophie anatomique, 1er vol. Méquignon-Marvis, Paris, France. Gerberding, M. 1997. Germ band formation and early neurogenesis of Leptodora kindti (Cladocera): first evidence for neuroblasts in the entomostracan crustaceans. Inv. Reprod. Dev. 32: 63–73. Gerberding, M. and G. Scholtz. 2001. Neurons and glia in the midline of the higher crustacean Orchestia cavimana are generated via an invariant cell lineage that comprises a median neuroblast and glial progenitors. Dev. Biol. 235: 397–409. Giribet, G. and S. Carranza, J. Baguna, M. Riutort, and C. Ribera. 1996. First molecular evidence for the existence of a Tardigrada + Arthropoda clade. Mol. Biol. Evol. 13: 76– 84. Giribet, G. and G.D. Edgecombe, and W.C. Wheeler. 2001. Arthropod phylogeny based on eight molecular loci and morphology. Nature 413: 157–161. Hallberg, E. and R. Elofsson. 1983. The larval compound eye of barnacles. J. Crust. Biol. 3: 17–24. Hanström, B. 1947. The brain, the sense organs and the incretory organs of the head in the Crustacea Malacostraca. Lunds Univ. Arsskrift. N.F. 43: 1–45. Harzsch, S. and J. Miller, J. Benton, R.R. Dawirs, and B. Beltz. 1998. Neurogenesis in the thoracic neuromeres of two crustaceans with different types of metamorphic development. J. Exp. Biol. 201: 2465–2479. Harzsch, S. 2001. Neurogenesis in the crustacean ventral nerve cord: homology of neuronal stem cells in Malacostraca and Branchiopoda? Evol. Dev. 3: 154–169. Harzsch, S. and D. Walossek. 2001. Neurogenesis in the developing visual system of the branchiopod crustacean Triops longicaudatus (LeConte, 1846): corresponding patterns of compound-eye formation in Crustacea and Insecta? Dev. Genes Evol. 211: 37–43. Harzsch, S. 2002. The phylogenetic significance of crustacean optic neurophils and chiasmata: a re-examination. J. Comp. Neurol. 453: 10–21. Harzsch, S. 2004. Phylogenetic comparison of serotonin-immunoreactive neurons in representatives of the Chilopoda, Diplopoda, and Chelicerata: implications for arthropod relationships. J. Morphol. 259: 198–213. Harzsch, S. and C.H. Muller, and H. Wolf. 2005. From variable to constant cell numbers: cellular characteristics of the arthropod nervous system argue against a sister-group relationship of Chelicerata and “Myriapoda” but favour the Mandibulata concept. Dev. Genes Evol. 215: 53–68. Harzsch, S. and G. Hafner. 2006. Evolution of eye development in arthropods: Phylogenetic aspects. Arthropod. Struct. Dev. 35: 319–340. Harzsch, S. and K. Vilpoux, D.C. Blackburn, D. Platchetzki, N.L. Brown, R. Melzer, K.E. Kempler, and B.A. Battelle. 2006. Evolution of arthropod visual systems: development of the eyes and central visual pathways in the horseshoe crab Limulus polyphemus Linnaeus, 1758 (Chelicerata, Xiphosura). Dev. Dyn. 235: 2641–2655.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

77

Hassanin, A. and N. Leger, and J. Deutsch. 2005. Evidence for multiple reversals of asymmetric mutational constraints during the evolution of the mitochondrial genome of metazoa, and consequences for phylogenetic inferences. Syst. Biol. 54: 277–298. Hassanin, A. 2006. Phylogeny of Arthropoda inferred from mitochondrial sequences: strategies for limiting the misleading effects of multiple changes in pattern and rates of substitution. Mol. Phylogenet. Evol. 38: 100–116. Hejnol, A. and G. Scholtz. 2004. Clonal analysis of Distal-less and engrailed expression patterns during early morphogenesis of uniramous and biramous crustacean limbs. Dev. Genes Evol. 214: 473–485. Hwang, U.W. and M. Friedrich, D. Tautz, C.J. Park, and W. Kim. 2001. Mitochondrial protein phylogeny joins myriapods with chelicerates. Nature 413: 154–157. Jager, M. and J. Murienne, C. Clabaut, J. Deutsch, H. Le Guyader, and M. Manuel. 2006. Homology of arthropod anterior appendages revealed by Hox gene expression in a sea spider. Nature 441: 506–508. Jenner, R.A. and G. Scholtz. Playing another round of metazoan phylogenetics: historical epistemology, sensitivity analysis, and the position of Arthropoda within the Metazoa on the basis of morphology. pp. 355–385. In: S. Koenemann and R.A. Jenner. [eds.] 2005. Crustacea and Arthropod Relationships. CRC Press, Boca Raton, USA. Jockusch, E.L. and C. Nulsen, S.J. Newfeld, and L.M. Nagy. 2000. Leg development in flies versus grasshoppers: differences in dpp expression do not lead to differences in the expression of downstream components of the leg patterning pathway. Development 127: 1617–1626. Jockusch, E.L. and T.A. Williams, and L.M. Nagy. 2004. The evolution of patterning of serially homologous appendages in insects. Dev. Genes Evol. 214: 324–338. Kadner, D. and A. Stollewerk. 2004. Neurogenesis in the chilopod Lithobius forficatus suggests more similarities to chelicerates than to insects. Dev. Genes Evol. 214: 367–379. Kraus, O. 2001. “Myriapoda” and the ancestry of the Hexapoda. Ann. Soc. Entomol. Fr. 37: 105–127. Kusche, K. and T. Burmester. 2001. Diplopod hemocyanin sequence and the phylogenetic position of the Myriapoda. Mol. Biol. Evol. 18: 1566–1573. Lankester, E.R. 1870. On the use of the term homology in modern zoology and the distinction between homogenetic and homoplastic agreements. Ann. Mag. Nat. Hist. Ser. 4. 6: 34– 43. Lankester, E.R. 1904. The structure and classification of the Arthropoda. Quart. J. Microscop. Sci. 47: 523–576. Lavrov, D.V. and W.M. Brown, and J.L. Boore. 2004. Phylogenetic position of the Pentastomida and (pan)crustacean relationships. Proc. Biol. Sci. 271: 537–544. Lewis, E.B. 1978. A gene complex controlling segmentation in Drosophila. Nature 276: 565– 570. Lim, J.T. and U.W. Hwang. 2006. The complete mitochondrial genome of Pollicipes mitella (Crustacea, Maxillopoda, Cirripedia): non-monophylies of maxillopoda and crustacea. Mol. Cells 22: 314–322. Loesel, R. and D.R. Nassel, and N.J. Strausfeld. 2002. Common design in a unique midline neurophil in the brains of arthropods. Arthropod. Struct. Dev. 31: 77–91. Mallatt, J.M. and J.R. Garey, and J.W. Shultz. 2004. Ecdysozoan phylogeny and Bayesian inference: first use of nearly complete 28S and 18S rRNA gene sequences to classify the arthropods and their kin. Mol. Phylogenet. Evol. 31: 178–191. Manuel, M. and M. Jager, J. Murienne, C. Clabaut, and H. Le Guyader. 2006. Hox genes in sea spiders (Pycnogonida) and the homology of arthropod head segments. Dev. Genes Evol. 216: 481–491. Maxmen, A. and W.E. Browne, M.Q. Martindale, and G. Giribet. 2005. Neuroanatomy of sea spiders implies an appendicular origin of the protocerebral segment. Nature 437: 1144–1148.

© 2011 by Taylor and Francis Group, LLC

78

Key Transitions in Animal Evolution

Meinertzhagen, I.A. and T.E. Hanson. The development of the optic lobe. pp. 1363–1491. In: M. Bate and A. Martinez Arias. [eds.] 1993. The Development of Drosophila melanogaster. CHCL Press, Cold Spring Harbor NY, USA. Melzer, R.R. and R. Diersch, D. Nicastro, and U. Smola. 1997. Compound eye evolution: Highly conserved retinula and cone cell patterns indicate a common origin of the insect and crustacean ommatidium. Naturwissenschaften 84: 542–544. Mittmann, B. 2002. Early neurogenesis of the horseshoe crab Limulus polyphemus and its implication for arthropod relationships. Biol. Bull. 203: 221–222. Mittmann, B. and G. Scholtz. 2003. Development of the nervous system in the “head” of Limulus polyphemus (Chelicerata: Xiphosura): morphological evidence for a correspondence between the segments of the chelicerae and of the (first) antennae of Mandibulata. Dev. Genes Evol. 213: 9–17. Morgan, T.H. 1928. The Theory of the Gene. Yale University Press, New Haven. Müller, C.H. and J. Rosenberg, S. Richter, and V.B. Meyer-Rochow. 2003. The compound eye of Scutigera coleoptrata (Linnaeus, 1758) (Chilopoda: Notostigmophora): an ultrastructural reinvestigation that adds support to the Mandibulata concept. Zoomorphology 122: 191–209. Negrisolo, E. and A. Minelli, and G. Valle. 2004. The mitochondrial genome of the house centipede Scutigera and the monophyly versus paraphyly of myriapods. Mol. Biol. Evol. 21: 770–780. Nielsen, C. 2001. Animal Evolution. Interrelationships of the Iiving Phyla. Oxford University Press, Oxford, UK. Nielsen, C. 2003. Proposing a solution to the Articulata-Ecdysozoa controversy. Zool. Script. 32: 475–482. Nilsson, D.-E. and D. Osorio. Homology and parallelism in arthropod sensory processing. pp. 333–347. In: R.A. Fortey and R.H. Thomas [eds.] 1997. Arthropod Relationships. Chapman & Hall, London. Nilsson, D.E. and A. Kelber. 2007. A functional analysis of compound eye evolution. Arthropod. Struct. Dev. 36: 373–385. Nulsen, C. and L.M. Nagy. 1999. The role of wingless in the development of multibranched crustacean limbs. Dev. Genes Evol. 209: 340–348. Nüsslein-Volhard, C. and E. Wieschaus. 1980. Mutations affecting segment number and polarity in Drosophila. Nature 287: 795–801. Oakley, T.H. and C.W. Cunningham. 2002. Molecular phylogenetic evidence for the independent evolutionary origin of an arthropod compound eye. Proc. Natl. Acad. Sci. USA 99: 1426–1430. Osorio, D. and J.P. Bacon. 1994. A good eye for arthropod evolution. BioEssays 16: 419–424. Osorio, D. and M. Averof, and J.P. Bacon. 1995. Arthropod evolution: great brains, beautiful bodies. Trends Ecol. Evol. 10: 449–454. Owen, R. 1849. On the Nature of Limbs. London John van Voorst, London, reprinted 2007, The Chicago University Press, Chicago.. Panganiban, G. and A. Sebring, L. Nagy, and S. Carroll. 1995. The development of crustacean limbs and the evolution of Arthropods. Science 270: 1363–1366. Panganiban, G. and S.M. Irvine, C. Lowe, H. Roehl, L.S. Corley, B. Sherbon, J.K. Grenier, J.F. Fallon, J. Kimble, M. Walker, et al. 1997. The origin and evolution of animal appendages. Proc. Natl. Acad. Sci. USA 94: 5162–5166. Park, S.J. and Y.S. Lee, and U.W. Hwang. 2007. The complete mitochondrial genome of the sea spider Achelia bituberculata (Pycnogonida, Ammotheidae): arthropod ground pattern of gene arrangement. BMC Genomics 8: 343. Paulus, H.F. 2000. Phylogeny of the Myriapoda—Crustacea—Insecta: a new attempt using photoreceptor structure. J. Zool. Syst. Evol. Res. 38: 189–208. Peterson, K.J. and D.J. Eernisse. 2001. Animal phylogeny and the ancestry of bilaterians: inferences from morphology and 18S rDNA gene sequences. Evol. Dev. 3: 170–205.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

79

Philippe, H. and N. Lartillot, and H. Brinkmann. 2005. Multigene analyses of bilaterian animals corroborate the monophyly of Ecdysozoa, Lophotrochozoa, and Protostomia. Mol. Biol. Evol. 22: 1246–1253. Pisani, D. 2004. Identifying and removing fast-evolving sites using compatibility analysis: an example from the Arthropoda. Syst. Biol. 53: 978–989. Pisani, D. and L.L. Poling, M. Lyons-Weiler, and S.B. Hedges. 2004. The colonization of land by animals: molecular phylogeny and divergence times among arthropods. BMC Biol. 2: 1. Popadic, A. 2005. Global evolution of nubbin expression patterns in arthropods: emerging view. Evol. Dev. 7: 359–361. Popadic, A. and G. Panganiban, D. Rusch, W.A. Shear, and T.C. Kaufman. 1998. Molecular evidence for the gnathobasic derivation of arthropod mandibles and for the appendicular origin of the labrum and other structures. Dev. Genes Evol. 208: 142–150. Popadic, A. and D. Rusch, M. Peterson, B.T. Rogers, and T.C. Kaufman. 1996. Origin of the arthropod mandible. Nature 380: 395. Prpic, N.M. and R. Janssen, B. Wigand, M. Klingler, and W.G. Damen. 2003. Gene expression in spider appendages reveals reversal of exd/hth spatial specificity, altered leg gap gene dynamics, and suggests divergent distal morphogen signaling. Dev Biol. 264: 119–140. Prpic, N.M. and D. Tautz. 2003. The expression of the proximodistal axis patterning genes Distal-less and dachshund in the appendages of Glomeris marginata (Myriapoda: Diplopoda) suggests a special role of these genes in patterning the head appendages. Dev. Biol. 260: 97–112. Prpic, N.M. 2004. Homologs of wingless and decapentaplegic display a complex and dynamic expression profile during appendage development in the millipede Glomeris marginata (Myriapoda: Diplopoda). Front Zool. 1: 6. Prpic, N.M. and W.G. Damen. 2004. Expression patterns of leg genes in the mouthparts of the spider Cupiennius salei (Chelicerata: Arachnida). Dev. Genes Evol. 214: 296–302. Prpic, N.M. and W.G.M. Damen. Arthropod appendages: a prime example for the eovlution of morphological diversity and innovation. pp. 381–398. In: A. Minelli and G. Fusco. [eds.] 2008. Evolving Pathways. Cambrige Univ. Press, Cambridge, UK. Prpic, N.M. and M.J. Telford. 2008. Expression of homothorax and extradenticle mRNA in the legs of the crustacean Parhyale hawaiensis: evidence for a reversal of gene expression regulation in the pancrustacean lineage. Dev. Genes Evol. 218: 333–339. Regier, J.C. and J.W. Shultz. 2001. Elongation factor-2: a useful gene for arthropod phylogenetics. Mol. Phylogenet. Evol. 20: 136–148. Regier, J.C. and H.M. Wilson, and J.W. Shultz. 2005. Phylogenetic analysis of Myriapoda using three nuclear protein-coding genes. Mol. Phylogenet. Evol. 34: 147–158. Richter, S. 2002. The Tetraconata concept: hexapod-crustacean relationships and the phylogeny of Crustacea. Org. Divers. Evol. 2: 217–237. Roeding, F. and S. Hagner-Holler, H. Ruhberg, I. Ebersberger, A. von Haeseler, M. Kube, R. Reinhardt, and T. Burmester. 2007. EST sequencing of Onychophora and phylogenomic analysis of Metazoa. Mol. Phylogenet. Evol. 45: 942–951. Ronco, M. and T. Uda, T. Mito, A. Minelli, S. Noji, and M. Klingler. 2008. Antenna and all gnathal appendages are similarly transformed by homothorax knock-down in the cricket Gryllus bimaculatus. Dev. Biol. 313: 80–92. Rota-Stabelli, O. and M.J. Telford. 2008. A multi-criterion approach for the selection of optimal outgroups in phylogeny: Recovering some support for Mandibulata over Myriochelata using mitogenomics. Mol. Phylogenet. Evol. 48: 103–111. Ryu, S.H. and J.M. Lee, K.H. Jang, E.H. Choi, S.J. Park, C.Y. Chang, W. Kim, and U.W. Hwang. 2007. Partial mitochondrial gene arrangements support a close relationship between Tardigrada and Arthropoda. Mol. Cells 24: 351–7. Schmidt-Rhaesa, A. 2001. Tardigrades: Are they really miniaturized dwarfs? Zool. Anz. 240: 549–555.

© 2011 by Taylor and Francis Group, LLC

80

Key Transitions in Animal Evolution

Schmidt-Rhaesa, A. and T. Bartolomeus, C. Lemburg, U. Ehlers, and J.R. Garey. 1998. The position of the Arthropoda in the phylogenetic system. J. Morphol. 238: 263–285. Scholtz, G. Cleavage, germ-band formation and head segmentation: the ground pattern of Euarthropoda. pp. 317–332. In: R.A. Fortey and R.H. Thomas. [eds.] 1997. Arthropod Relationships. Chapman & Hall, London. Scholtz, G. and B. Mittmann, and M. Gerberding. 1998. The pattern of Distal-less expression in the mouthparts of crustaceans, myriapods and insects: new evidence for a gnathobasic mandible and the common origin of Mandibulata. Int. J. Dev. Biol. 42: 801–810. Scholtz, G. 2001. Evolution of developmental patterns in arthropods—the analysis of gene expression and its bearing on morphology and phylogenetics. Zoology 103: 99–111. Scholtz, G. 2002. The Articulata hypothesis—or what is a segment? Org. Divers. Evol. 2: 197–215. Scholtz, G. Is the taxon Articulata obsolete? Arguments in favour of a close relationship between annelids and arthropods. pp. 489–501. In: A. Legakis, S. Sfenthorakis, R. Polymeni and M. Thessalou-Legaki. [eds.] 2003. The New Panorama of Animal Evolution. Pensoft, Sofia-Moscow. Scholtz, G. 2005. Homology and ontogeny: Pattern and process in comparative developmental biology. Theor. Biosci. 124: 121–143. Scholtz, G. and G.D. Edgecombe. 2006. The evolution of arthropod heads: reconciling morphological, developmental and palaeontological evidence. Dev. Genes Evol. 216: 395–415. Sewell, W. and T. Williams, J. Cooley, M. Terry, R. Ho, and L. Nagy. 2008. Evidence for a novel role for dachshund in patterning the proximal arthropod leg. Dev. Genes Evol. 218: 293–305. Shear, W.A. 1992. End of the ‘Uniramia’ taxon. Nature 359: 477–478. Shultz, J.W. and J.C. Regier. 2000. Phylogenetic analysis of arthropods using two nuclear protein-encoding genes supports a crustacean + hexapod clade. Proc. Biol. Sci. 267: 1011–1019. Sinakevitch, I. and J.K. Douglass, G. Scholtz, R. Loesel, and N.J. Strausfeld. 2003. Conserved and convergent organization in the optic lobes of insects and isopods, with reference to other crustacean taxa. J. Comp. Neurol. 467: 150–172. Snodgrass, R.E. 1935. Principles of Insect Morphology. McGraw-Hill, New York, USA. Stollewerk, A. 2002. Recruitment of cell groups through Delta/Notch signalling during spider neurogenesis. Development 129: 5339–5348. Stollewerk, A. and D. Tautz, and M. Weller. 2003. Neurogenesis in the spider: new insights from comparative analysis of morphological processes and gene expression patterns. Arthropod. Struct. Dev. 32: 5–16. Stollewerk, A. Evolution of neurogenesis in arthropods. pp. 359–380. In: A. Minelli and G. Fusco [eds.] 2008. Evolving pathways. Cambrige Univ. Press, Cambridge, UK. Stollewerk, A. and A.D. Chipman. 2006. Neurogenesis in myriapods and chelicerates and its importance for understanding arthropod relationships. Integr. Comp. Biol. 46: 195– 206. Stollewerk, A. and P. Simpson. 2005. Evolution of early development of the nervous system: a comparison between arthropods. Bioessays 27: 874–883. Strausfeld, N.J. 1998. Crustacean-insect relationships: the use of brain characters to derive phylogeny amongst segmented invertebrates. Brain Behav. Evol. 52: 186–206. Strausfeld, N.J. 2005. The evolution of crustacean and insect optic lobes and the origins of chiasmata. Arthropod. Struct. Dev. 34: 235–256. Telford, M.J. and R.H. Thomas. 1995. Systematics: Demise of the Atelocerata? Nature 376: 123–124. Telford, M.J. and R.H. Thomas. 1998. Expression of homeobox genes shows chelicerate arthropods retain their deutocerebral segment. Proc. Natl. Acad. Sci. USA 95: 10671– 10675.

© 2011 by Taylor and Francis Group, LLC

Pending Issues in Development and Phylogeny of Arthropods

81

Telford, M.J. and S.J. Bourlat, A. Economou, D. Papillon, and O. Rota-Stabelli. 2008. The evolution of the Ecdysozoa. Philos. Trans. R. Soc. Lond. B Biol. Sci. 11: 1529–1537. Timmermans, M.J.T.N. and D. Roelofs, J. Marien, and N.M. van Straalen. 2008. Revealing pancrustacean relationships: Phylogenetic analysis of ribosomal protein genes places Collembola (springtails) in a monophyletic Hexapoda and reinforces the discrepancy between mitochondrial and nuclear DNA markers. BMC Evol. Biol. 8: 83. Turbeville, J.M. and D.M. Pfeifer, K.G. Field, and R.A. Raff. 1991. The phylognetic status of Arthropods, as inferred from 18S rRNA sequences. Mol. Biol. Evol. 8: 669–686. Ungerer, P. and G. Scholtz. 2008. Filling the gap between identified neuroblasts and neurons in crustaceans adds new support for Tetraconata. Proc. Biol. Sci. 275: 369–376. Wegerhoff, R. and O. Breidbach. Comparative aspects of the chelicerate nervous system. pp. 159–179. In: O. Breidbach and W. Kutsch. [eds.] 1995. The Nervous System of Invertebrates: An Evolutionary and Comparative Approach. Birkhauser, Basel. Whitington, P.M. Conservation versus change in early axogenesis in arthropod embryos: A comparison between myriapods, crustaceans and insects. pp. 181–219. In: O. Breidbach and W. Kutsch [eds.] 1995. The Nervous System of Invertebrates: An Evolutionary and Comparative Approach. Birkhauser, Basel. Whitington, P.M. 1996. Evolution of neural development in arthropods. Sem. Cell Dev. Biol. 7: 605–614. Williams, D.S. and P. McIntyre. 1980. The principal eyes of a jumping spider have a telephoto component. Nature 288: 578–580. Williams, T.A. 1998. Distal-less expression in crustaceans and the patterning of branched limbs. Dev. Genes Evol. 207: 427–434. Williams, T.A. 1999. Morphogenesis and homology in arthropods limbs. Amer. Zool. 39: 664–675. Williams, T.A. and C. Nulsen, and L.M. Nagy. 2002. A complex role for Distal-less in crustacean appendage development. Dev. Biol. 241: 302–312. Williams, T.A. 2008. Early Distal-less expression in a developing crustacean limb bud becomes restricted to setal-forming cells. Evol. Dev. 10: 114–120. Wilson, K. and V. Cahill, E. Ballment, and J. Benzie. 2000. The complete sequence of the mitochondrial genome of the crustacean Penaeus monodon: are malacostracan crustaceans more closely related to insects than to branchiopods? Mol. Biol. Evol. 17: 863–874. Zantke, J. and K. Wolff, and G. Scholtz. 2008. Three-dimensional reconstruction of the central nervous system of Macrobiotus hulefandi (Eutardigrada, Parchela): implications for the phylogenetic position of Tardigrada. Zoomorphology 127: 21–36. Zrzavy, J. and V. Hypsa, and M. Vlaskova. Arthropod phylogeny: taxonomic congruence, total evidence and conditional combination approaches to morphological and molecular data sets. pp. 97–107. In: R.A. Fortey and R.H. Thomas [eds.] 1997. Arthropod Relationships. Chapman & Hall, London, UK.

© 2011 by Taylor and Francis Group, LLC

Section 2 The Earliest Animals: From Genes to Transitions

© 2011 by Taylor and Francis Group, LLC

Chapter 5

The Pre-Nervous System and Beyond—Poriferan Milestones in the Early Evolution of the Metazoan Nervous System* Michael Nickel

Introduction Metazoan nervous systems are structurally and functionally diverse and highly complex. Even the simplest nervous systems, the nerve nets of Cnidaria and Ctenophora, showcase a structural complexity on the cellular and sub-cellular level which implies a long evolutionary history. The nervous system in a strict sense, as defined by most authors (for overviews see Bullock and Horridge 1969, Hanström 1928, Schmidt-Rhaesa 2007), is an autapomorphy of the Eumetazoa. One of the definitions is provided by Schmidt-Rhaesa (2007): “Nervous systems conduct information in a directed way through the body. This is done by electrical and/or chemical signals and by cells specialized for these functions”. The classic neurone doctrine by Ramon y Cayal (1937), however, is strictly morphology-based and does not consider the nature of the signal. Bullock and Horridge (1969) summarized Ramon y Cayals’s doctrine by stating that “all nervous systems consist in essence (whatever other, nonnervous elements may be Institut für Spezielle Zoologie und Evolutionsbiologie Friedrich-Schiller-Universität Jena, Erbertstr. 1, D-07743 Jena Germany. E-mail: [email protected] *This chapter is dedicated to Stephen Hillenburg, who greatly directed the interest of a whole future scientist generation towards sponges and the marine life.

© 2011 by Taylor and Francis Group, LLC

86

Key Transitions in Animal Evolution

present) of distinct cells called neurons, which are specialized for nervous functions and which produce prolongations and branches”. As we will see, this very formal definition resulted in an academic dispute if there is a nervous system in sponges. This question was negated based on solid arguments (Jones 1962), however it tends to reappear in literature until today (e.g. Meech and Mackie 2007, Perovic et al. 1999). The phylogenetic context of the early evolution of the nervous system is difficult to consider, since sponges (Porifera) and Placozoa do not display distinct nervous system structures, namely defined nerve cells and synapses. For this reason, the evolution of the central nervous systems (CNS) of the Bilateria acquires the most attention (Bullock and Horridge 1969, Hanström 1928, Schmidt-Rhaesa 2007). However the evolutionary tendency for centralization is already intrinsic within the Cnidaria (Holland 2003). Nonetheless, ‘simple’ nerve nets and the properties of their neurons are generally the focus of literature on early nervous system evolution (Mackie 1970, Meech and Mackie 2007, Pantin 1952, 1956, Parker 1910, 1919, Passano 1963). This comprehension of ‘early nervous system evolution’ is synonymous with ‘early evolution of electrical excitability’. If we want to understand the basics, we have to consider how the first metazoa, the LCAM (last common ancestor of metazoa), sensed external stimuli, how it facilitated intercellular signaling and integration. Therefore, we will have to reflect on the evolution of pre-nervous integration systems. Even though sponges lack a nervous system, poriferan milestones of nervous system evolution will provide decent information in helping to reconstruct the LCAM’s integrative capabilities and possible pre-nervous system intermediates. The reason, why sponges are important in this context is simple: they display movements without muscles and signal integration without nerves (Nickel 2007). Their coordinated movements are extremely slow: contractions often follow rhythmic patterns and are easier to recognize by the human eye than the extremely slow locomotion of the body (Nickel 2004, 2006). The first reports of these behaviors, dating back to Aristotle, gave rise to one of the longest scientific debates in the history of science, the question if sponges are animal or plants or something in-between. In addition, this behavior and the underlying biological mechanisms and principles are the key to our main questions on how integration systems evolved from the LCAM to the Cnidaria. Here, I aim to provide an overview of the current knowledge on sponge contraction, locomotion and integration, which will provide first evidence for the early co-evolution of contraction and integration in Metazoa. From this background, I will deduce a model for a pre-nervous system of the demosponges and I will develop a hypothesis on the evolutionary steps from the pre-nervous to the nervous system. The chapter will start with an

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

87

overview on the long history of research on contraction and integration. This historical view will facilitate the understanding of present concepts, new hypotheses and the needs for future research. Even though I will correct some historical miss-interpretations (e.g. Jones 1962, concerning Aristotle’s views on sponges), readers who are well aware about the historical context might want to continue directly with the paragraphs on current concepts and new conclusions below.

The History of Research on Contractile and Integrative Systems in Sponges Δοκεῖ δὲ καὶ ὁ σπόγγος ἔχειν τινὰ αἴσθησιν1 Aristotle 350 B.C. (from Barthélemy-Saint Hilaire 1883) Contraction: A Hint from the Classical Antiquity The fact that sponges were for long periods regarded as plants (Linné 1767) or “Pflanzenthiere” (Esper 1794–1805) points out the difficulties of categorizing sponges. Generations of biologists seem to have had difficulties in how to interpret sponges on the scale from plant to animals (for an early review see Johnston 1842). One must not however take this as a sign of the superiority of present day science or scientific advancement over the centuries. It is likely more likely of a sign for varying perceptive concepts in biological science over the centuries. In fact, going back to the first written reports on sponges, we find that sponges were recognized as animals very early on. As early as 350 B.C., in his “Natural History of Animals”, Aristotle (384–322 B.C.) addresses sponges as animals (Aristotle 1498). Unfortunately, this historically important fact is not generally recognized by zoologists (e.g. Meech 2008 argues that Aristotle regarded sponges as vegetables, which is only valid for parts of Aristotle’s manuscripts). Aristotle mentions sponges in his first chapter of book one, characterizing them as sedentary marine animals, and attributing sensibility to them. This sensibility is expressed by the fact that, when not done carefully, they are hard to remove from the substrate. From chapter 16 of his book five (Fig. 1), which almost exclusively deals with sponges, it is evident that he is referring to sponge contraction. Aristotle reiterates that sponges are sensitive and he relates contractile behavior to sponge fishing and environmental conditions. He states that they are able to sense when they are about to be collected by sponge fisherman, and how they 1 “And, by the way, the sponge appears to be endowed with a certain sensibility”; Aristotle, 1910. The History of Animals. Transl. by D’Arcy Wentworth Thompson. Oxford: Clarendon Press. [Electronic reprint]. The University of Adelaide Library. http://etext.library.adelaide. edu.au/a/aristotle/history/.

© 2011 by Taylor and Francis Group, LLC

88

Key Transitions in Animal Evolution

Fig. 1. Facsimile of Aristotle’s History of the animals in the Latin translation, printed in 1498, page from book 5, chapter 16 which exclusively deals with sponges. The highlighted passage reads: “It is said that the sponge is sensitive; and as a proof of this statement they say that if the sponge is made aware of an attempt being made to pluck it from its place of attachment it draws itself together, and it becomes a difficult task to detach it. It makes a similar contractile movement in windy and boisterous weather, obviously with the object of tightening its hold. Some persons express doubts as to the truth of this assertion; as, for instance, the people of Torone.” (Aristotle, 1910; translation by D’Arcy Wentworth Thompson). For almost 2200 years, between its genesis 350 B.C. to around 1850, ‘Aristotle’s History’ served as an integral source of zoological knowledge and was regularly cited (compare Johnston, 1842). Modified scan from Aristotle (1498).

react to this with contraction. This results in increasing efforts for the fishermen to remove them from the substrate. The same behavior can also be found during turbulent seas, when adhesion is even more enhanced. Interestingly, he adds that this behavior is questioned by the fishermen of Toroni; who presumably, did not share this opinion (Fig. 1). These short passages by Aristotle are interesting in multiple ways, for science history as well as for zoology in general. First, he regards sponges as animals, a classification which was later refused until the mid 18th century, when Ellis (1755), Pallas (1766) as well as Ellis and Solander (1786)

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

89

reestablished this view. Second, he describes sponge contractile behavior, which doesn´t reappears in scientific literature before Pallas (1787). In addition, Aristotle interprets this behavior as a clear sign of integration of information in the sponge. The dissent of the Toroni fishermen he mentions may be seen as a first sign of a much later scientific debate on the sponge nervous system (see below). Similar reports on sponges can be found in texts of various authors of the classical antique. For example, Claudius Aelianus (approx. 175–235 A.C.) mentions very similar aspects in his book “De Natura Animalium” (book 8, chapter 16). The context suggests that his is a review of Aristotle’s text, rather than an independent report (Aelianus 1832). More than a century earlier, Plinius the older (23–79 A.C.) reported on sponges’ contractile behavior as well as their ability to regenerate from their “roots” (Plinus 1855). Sponge body contractions: Perception in modern biology The earliest modern report on sponge contraction seems to be the one of Pallas (1787). Most likely inspired by the three antique documents mentioned above, he investigated sponge behavior and observed oscular contraction. Also the slightly earlier work of Ellis and Solander (1786) mentions Aristotle’s description of sponge contraction, although they did not add their own observations to the scientific record. Their work demonstrates par excellence that the knowledge of the antique natural history influenced natural sciences at least until the mid 18th century. It took some more decades, before zoologists started to examine the details of sponge contractions themselves on a broader basis. As we learn from Johnston (1842), sponge contractility played a certain role in the academic discussion on the animal or plant nature of sponges until the late 18th century. However, he himself stated in his nicely detailed review: “Sponges are unmoving and unirritable: hence they ever remain rooted to the place of their germination, and are incapable either of contracting or dilating themselves, or even of moving any fibre or portion of their mass” (Johnston 1842). Only some years later, Bowerbank (1856) presented his extensive long-time observations of oscules opening and closing as well as surface contractions. Lieberkühn (1859) reported the contractility of the body surface (exopinacoderm) of Spongia tupha PALLAS 1766. A few years later, Schmidt was the first to postulate a cellular mechanism for contraction. He suggested contractile, muscle like fibers, which he saw as the “erste wohl constatirte Beispiel des Vorkommens (willkürlich) contractiler Fasern im Körper der Spongien” [first well documented example of the existence of (voluntarily) contractile fibers in the body of the sponges]. In relation to Tethya sp. he spoke of the presence of muscle fibers within the cortex (Schmidt 1866).

© 2011 by Taylor and Francis Group, LLC

90

Key Transitions in Animal Evolution

One of the first graphical representations of a contracted sponge was published by the end of the 19th century on the zoological wall charts edited by Rudolf Leuckart (see Redi et al. 2002). Drawn and described by Marshall (1885), chart 47 compared a contracted and expanded specimen of Tethya sp. in cross sections. His remarkable drawings show the contraction of the cortex as well as parts of the aquiferous system (Fig. 2). The first detailed drawing of the contraction of a calcareous sponge was presented by Minchin (1900). He demonstrated the contraction of the pinacoderm as well as the resulting displacement of spicules. Muscle fiber cells, or rather spindle shaped myocytes were postulated by Schmidt (1866), Sollas (1888) and other authors (for an overview see Jones 1962). Whereas Minchin (1900) and Wilson (1910) observed pinacoderm contraction. Consequently, two competing hypotheses on the

Fig. 2. Early published drawings by Marshalls (1885) showing for the first time morphological changes during contraction in Tethya sp. Collage of two drawings present on Leuckart’s zoological wall chart 47. Below, the detailed explanatory note on wall chart figure 3 (contraction, upper right) originally printed in a leaflet accompanying the wall chart is presented (combined and modified from Redi 2002).

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

91

contractile effectors emerged: (1) mesohyle-mediated contraction based on myocytes (preferable terminus after Boury-Esnault and Rützler 1997: actinocytes); (2) pinacoderm-mediated contraction based on pinacocytes. Most of the later authors preferred the first hypothesis. Among the more recent works, Bagby (1966) first pointed out the similarities between the spindle-shaped myocytes and smooth muscle cells, partly because of the staining characteristics using Mallory’s trichrome. In addition, he ultrastructurally demonstrated the presence of actin filament bundles (Bagby 1966). Several years later, however, he found similar filament structures in pinacocytes (Bagby 1970), which was confirmed in subsequent studies (Matsuno et al. 1988). Also in the 1960’s, Pavans de Cecatty became one of the main supporters of the myocyte hypothesis, connecting it to the discussion on the existence of a nervous system in sponges (see below: The Quest for the Elementary Nervous System). Mainly based on histological and ultrastructural evidence, e.g. the demonstration of cell-cell contacts and synapse-like structures, he defined myocytes as evolutionary prototypes of neuromuscular cells (Pavans de Ceccatty 1960, 1971, 1974a, 1979). Myocytes were found to be present in a number of sponges and accepted as contractile effectors by most authors (Lentz 1966, 1968, Prosser et al. 1962, Vacelet 1966). Contemporaneously, Jones (1962) pointed out missing functional proof-of-concepts regarding myocytes contractility. He referred to a number of publications which demonstrated contractility of pinacocytes directly or indirectly (Bidder 1937, Jones 1957, Parker 1910, Wilson 1910). Finally, after some years of discussion, Pavans de Cecatty integrated contractile pinacocytes into his neuromuscular model and demonstrated the presence of actin filaments in pinacocytes (Pavans de Ceccatty 1981, 1986). Nevertheless, he did not change his view about myocytes as the main contractile effectors in sponges. Simpson (1984) compiled ‘The Cell Biology of Sponges’ and provided a unique secondary literature source. His book has had the highest possible impact on the sponge research community and is still a major work of reference. However, it seems that during the two decades following publication of his book, the quantity of publications on sponge cell biology decreased, apart from some exceptions (e.g. Weissenfels 1989). This might be a coincidence or a result of the surpassing richness in detail provided by Simpson. His work also might have provoked the assumption that most about sponge biology is already known. Interestingly, only few works on sponge contraction were published from 1984 to 2004 (Matsuno et al. 1988, Sarà et al. 2001, Weissenfels 1990). However, after 20 years of relative silence, a new interest in the topic has arisen (Elliott and Leys 2007, Meech 2008, Nickel 2004, Nickel et al. 2006a, Nickel et al. 2006b, Nickel et al. 2005; for details see below: Current Concepts on Sponge Contraction, Locomotion and Integration).

© 2011 by Taylor and Francis Group, LLC

92

Key Transitions in Animal Evolution

The Late discovery of sponge locomotion The second behavioral aspect in sponges is locomotion. This was hidden from zoologists for a long time and is even today hard to find in zoology textbooks, although some notable exceptions should be mentioned (Ruppert et al. 2004, Van Soest 1996). Locomotion, however, is not the most obvious behavior, since poriferans are sessile animals. Almost all species are firmly attached to the substrate, with only a few species drifting passively. Nevertheless, many, if not all, sponge species display movements on the substrate. These movements are too slow to be observed by the human eye without the aid of the one or the other timelapse techniques. The most simple techniques are drawing outlines (e.g. Arndt 1941, Bond and Harris 1988), but today digital time-lapse imaging is used (Nickel 2006). However, understanding sponge locomotion, even more than understanding sponge contraction, requires the adoption of a time philosophy and a time scale which differs significantly from all other animals. The first published report on sponge locomotion seems to date back to Carter (1848). He recognized the slow displacement of adult freshwater sponges on the walls of water tanks. Lieberkühn (1863) followed with his observation of spongillid fragments moving apart from each other. Noll (1881) reported on the movement of a specimen of Reniera sp. growing on an oyster shell in his aquarium. Similar aquarium based observations were also reported by Arndt (1941), followed by Burton (1948), who included field observations. Most authors point out that shape changes are part or consequence of the locomotion sequences, suggesting the influence of differential growth (Arndt 1941). In contrast, very prominent movements which are not correlated with body shape changes were reported by Fishelson (1981). Individuals of Tethya seychelensis (WRIGHT 1841) and T. aurantium (PALLAS 1766) sized 20–30 mm moved 50–80 mm within 24 hours. Fishelson postulated that traction forces are generated by myocytes within the filament-like body extensions. He concludes that the extensions which are able to anchor to the substrate mediate physical forces between the sponge and its physical environment. Kilian (1967) observed negative phototaxis for juvenile freshwater sponges (Ephydatia spp. and Spongilla spp.) after hatching from gemmules. In contrast, Heteromeyena sp. displayed positive phototaxis. The strength of phototactic behavior is positively correlated with concentration of associated green algae within the sponges. These movements might be explained by amoeboid movements of the basal pinacoderm, shifting the whole sponge (Brøndsted 1942). Several studies investigated the movements of sponge masses after hatching from gemmules or of sponge reaggregates after artificial dissociation (for an overview see Jones 1962,

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

93

Wintermann 1951). Most of these studies confirmed amoeboid movements. However, one has to be aware that in all animals morphogenetic processes are associated with amoeboid cell movements and body shape changes. This is not necessarily equivalent to the movements observed in adult sponges shown above. As mentioned before, during the late 1980s interest in sponge cell biology seemed to declined (see: Sponge Body Contractions: Perception in Modern Biology), while the interest in sponge locomotion seemed to hold the same level. The reason might be found in the wider availability of analog and digital time-lapse imaging methods, which are used by recent researchers (Bond 1992, Bond and Harris 1988, Nickel 2006, Nickel and Brümmer 2002, Nickel and Brümmer 2004, Nickel et al. 2002, Pronzato 2004). The Quest for the Elementary Nervous System The question of an integrative system in sponges is closely related to the phenomena of contraction and locomotion. Consequently, questions and assumptions of a possible nervous system in sponges came up early. As outlined above, antique authors regarded sponges as ‘sensitive’. However, these questions did not only arise in sponges, but also in other animal groups. Based on the growing knowledge of the cnidarian nervous systems at the beginning of the 20th century, the first models on the evolution of nervous systems and ‘the elementary nervous system’ were published. Starting with Parker (1910, 1919), a century of general publications on the ‘primitive nervous system’ followed, indicated by titles combined from words like ‘sponges’, ‘integration’, ‘conduction’ and ‘coordination’: Pantin (1952, 1956), Jones (1962), Passano (1963), Lentz (1968), Pavans de Cecatty (1974a, b, 1979, 1989), Mackie (1970, 1979, 1990), and most recently Leys and Meech (2006). Based on experiments using cnidarians and poriferans, Parker tried to reconstruct the ‘elementary nervous system’: the evolutionary initial nervous system which structurally and functionally represents all elements of a nervous system in their most simple state. As a prerequisite, ‘the nervous system’ and its elements need to be fundamentally defined. This happened to be much easier in Parker’s lifetime than today, as Mackie (1990) points out. The enormously grown and growing biological knowledge makes it difficult to compile a list of characters defining a nerve cell. This refers especially to the knowledge on the identity of genetic information in every cell of a single individual as well as differentiation mechanisms via regulated gene expression (Anderson 1990). This is caused by the fact that many so-called ‘nerve cell typical’ features and molecules play a role in other biological contexts. Prominent examples are voltage-gated ion channels or transmitter substances (Mackie 1990).

© 2011 by Taylor and Francis Group, LLC

94

Key Transitions in Animal Evolution

According to Parker (1919) the first nervous system evolved from biological necessity to control ‘independent effectors’, namely contractile cells. Consequently, muscle cells are the evolutionary nucleus, around which the nervous system evolved. From his own studies on Stylotella sp. he concluded that sponges possess independent contractile effectors (pinacocytes and porocytes). In addition, he demonstrated the transmission of a mechanical stimulus, which after several minutes caused the contraction of an oscule several millimeters away. He spoke about ‘neuroid’ signal transduction, which ‘creeps’ through the sponge tissue, and he interpreted this observation as a form of transduction within the effector tissue itself. Therefore, he clearly distinguished it from his concept of signal transduction through the neuronal reflex arc, which connects sensor cells or pacemaker cells and ‘dependent effectors’ (e.g. epithel-muscle cells in Cnidaria) via nerve cells. This concept will have to be discussed below in the context of new molecular and physiological results. McNair (1923) reported similar results on the mechanical stimulation of contraction and the transmission through sponge tissue in Ephydatia fluviatilis (LINNAEUS 1759). The contractile wave traveled with a speed of 35 µm·sec–1 (downwards the oscule) and 17 µm·sec–1 (upwards the oscule). Similar speeds of contractile waves ranging from 8 to 30 µm·sec–1 were reported for E. fluviatilis, Spongia sp. and Hippospongia sp. by other authors (De Vos and Van De Vyver 1981, Pavans de Ceccatty 1971, 1974a). Jones (1962) tried to explain the directional differences in speed of the contractile waves observed by McNair (1923) by attributing upwards movements to the endopinacoderm and downwards movement to the exopinacoderm which are structurally different. He justified his view by referring to a report by Ankel (1948), who demonstrated structural differences between the two pinacoderm layers: exopinacocytes are oriented lengthwise in relation to the oscule, while endopinacocytes are oriented crosswise. Consequently, Jones argued, the signal crosses more cell borders inside the endopinacoderm. However, he did not explain the possible mechanisms limiting a signal to travel in one direction within a certain cell layer. In this context, his suggestion seems to be not very likely. In their reviews, Pantin (1952, 1956) and Passano (1963) modified the hypothesis of Parker, mainly based on their work in cnidarians. Consequently, their hypotheses on the evolution of nervous systems focused on electrical conduction, like Jones (1962) did. Nevertheless, all of them admitted to an evolutionary precursor system for ‘neuroid’ conduction. Macke (1970) argued that electrical synapses (gap junctions) likely evolved before chemical synapses and are therefore candidates for a neuroid pre-nervous system. This view was supported by the report of electrical coupling of Haliclona sp. cells by Loewenstein (1967). Both works resulted in an intensified search for gap junctions in sponges. Admittedly,

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

95

two independent groups found ultrastructural evidence for gap junctions (Gaino and Sarà 1976, Green and Bergquist 1979). However, this could not be unequivocally confirmed by the same and other authors (Garrone and Lethias 1990, Garrone et al. 1980, Green and Bergquist 1982, Lethias et al. 1983). No further work focusing on gap junctions in sponges has been published after 1990. Until today, no direct electrical conduction has been shown in demosponges and calcareous sponges. This might be an effect of the small cell sizes (Leys and Meech 2006, Mackie 1990). However, action potentials pass on through the trabecular tissue of hexactinellid sponges and influence ciliar beating of choanocytes (Leys and Mackie 1997, Leys and Mackie 1999, Leys et al. 1999, Mackie et al. 1983). Nevertheless, due to the syncytical nature of the trabecular system in hexactinellids (Leys et al. 2007), no nervous system like structures (e.g. synapses) are necessary to distribute the action potential. Since ciliates produce action potentials, as well, it is evident that voltage-gated ion channels, namely voltage-gated Ca2+-channels (Linder and Schultz 2002, Schultz et al. 1986), predate the evolution of animal multicellularity. An alternative scenario for the early evolution of the nervous system emphasizes the role of secretion (Grundfest 1959, 1965, Horridge 1968). Lentz (1966, 1968) demonstrated histochemically acetylcholine esterase activity as well as the presence of adrenalin and serotonin in the calcareous sponge Sycon sp. Activity was mainly located to actinocytes, and granulated as well as multipolar cells. In demosponges, Acetylcholine esterase activity was also demonstrated: in Hippospongia sp. (Thiney 1972) as well as in Tethya wilhelma SARÀ, SARÀ, NICKEL & BRÜMMER 2001 (Nickel 2001). In physiological experiments, acetylcholine influenced the rhythm as well as the intensity of oscule contractions of Spongia officinalis LINNAEUS 1759 (Pavans de Ceccatty 1971). Evidence presented by Emson (1966) on the effect of the acetylcholine receptor antagonist nicotine upon the choanocytes of Cliona celata GRANT 1826 is questionable, since non-physiological concentrations were used, which might have causeed cytotoxic effects. However, in the same work Emson (1966) showed the influence of adrenaline on the pumping behavior of C. celata, while Pavans de Cecatty (1971) found an increased contractile behavior in S. officinalis. Weyrer et al. (Weyrer et al. 1999b) established serotonin in the larvae of Tedania ignis (DUCHASSAING & MICHELOTTI 1864). These scattered demonstrations of neuroactive substance effects in sponges was completed by the cloning of a metabotrophic GABA/glutamate receptor in Suberites domuncula (OLIVI 1792) and the demonstration of GABA effecting intracellular Ca2+ concentrations (Perovic et al. 1999). Quite in parallel to the research situation on cell biology (see above: Sponge Body Contractions), the research interest on integration in sponges

© 2011 by Taylor and Francis Group, LLC

96

Key Transitions in Animal Evolution

seems to have declined between the 1980s and the dawn of the 21st century. However, at least several groups have kept up discussions on the topic (Mackie 1990, Pavans de Ceccatty 1989). In addition, during that period, Leys and co-workers demonstrated intracellular electrical potential changes and action potentials in the trabecular tissue of hexactinellid sponges (Leys and Mackie 1997, Leys and Mackie 1999, Leys et al. 1999, Mackie et al. 1983). These results provide clear evidence for the presence of voltage-gated ion channels in sponges, at least the Hexactinellida, as shown above. Taking into account the complete historic sequence of discussions about the evolution of the first nervous systems it becomes evident that they lead to a number of misunderstandings. This seems to be quite obvious, as shown by the more or less open verbal attacks amongst exponents of the various hypotheses between the 1960’s and the 1980’s. Despite of the polarization, it has to be emphasized that the work of Jones (1962) is still the most comprehensive work of the publications. Some of his comments and conclusions, however, are now outdated. Notably, Mackie and Pavans de Ceccatty attempt a constructive debate of the topic, especially in their reviews (Mackie 1970, 1979, 1990, Pavans de Ceccatty 1974a, b, 1979, 1989). Their intention was not to manifest a nervous system in sponges. This has sometimes been misinterpreted by other authors. Mackie, whose own work mainly reflected aspects of the nervous systems of cnidarians, emphasized repeatedly that the understanding of integration in sponges is the key to the understanding of the evolution of the first nervous systems (Mackie 1979, 1990), which seems to contrast the opinion of Jones. According to Mackie, the pharmacological and histological evidence supports that the same mechanisms favoring the evolution of nervous systems in other metazoans acted in sponges. In this sense, sponges represent a prime example on the mechanistic possibility of typical metazoan features like the integrated mechanical response upon external stimuli without nerves or muscles (Mackie 1990). Pavans de Ceccatty supported similar views. His comprising neuromuscular model for sponges includes hard-wired inter-cellular signaling in combination with permanently morphologically re-modulating signaling pathways (Pavans de Ceccatty 1974a). This model is still up-to-date and is neither proven in all details nor refuted. The recent review (Leys et al. 2007) and reiteration of some new evidence (Meech 2008) has demonstrated the actuality and the interest on the topic. Recent physiological and molecular evidence and further ongoing work will be needed to provide more detailed evidence to validate or disprove the historic views.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

97

Current Concepts on Sponge Contraction, Locomotion and Integration Sponge contraction and locomotion Phylogenetic context In which sponge groups do we find locomotion and contraction? In a phylogenetic context, contraction and locomotion are unevenly distributed among the recognized sponge clades. Within all sponge groups, contraction seems to be distributed wider than locomotion. Contraction is frequently found in Demospongiae (e.g. Arndt 1941, Marshall 1885, Nickel 2004, Schmidt 1866, for an example see Fig. 3a), Homoscleromorpha (unpublished data on Oscarella lobularis (Schmidt 1862), Fig. 3b) and Calcispongia (Minchin 1900, and see 3c). No contraction has been reported for Hexactinellida to date, which is not surprising, considering their rigid skeletons and their relatively low amount of cellular mass and extracellular matrix in relation to the silica skeleton (Leys et al. 2007). However, the same can be said about a number of Demospongiae clades. Those with rigid, spicule rich skeletons, like Petrosiidae VAN SOEST 1980 or Suberitidae SCHMIDT 1870 do not display significant external body contractions. Nevertheless, most of the species, if not all in these groups, will show oscular contractions (unpublished observations during SCUBA diving). In an x-ray microtomography based comparison of a glutamate treated specimen of Suberites domuncula and a control specimen, the treated specimen displayed internal contraction of

Fig. 3. Contraction in sponges. Line up of expanded (first column) and contracted states (second column) of sponge individuals and an overlay of both states (third column). Scale bars represent 5 mm. A. Demospongiae: Tethya wilhelma (see Nickel 2004). B. Homoscleromorpha: Oscarella lobularis (Nickel 2010). C. Calcispongia: Clathrina clathrus (Nickel 2010).

© 2011 by Taylor and Francis Group, LLC

98

Key Transitions in Animal Evolution

the aquiferous system (unpublished data). This situation is very similar to internal contractions in sponge species without such massive, rigid skeletons, which display external body contractions, e.g. Ephydatia mülleri (LIEBERKÜHN 1855) (Elliott and Leys 2007). Whether Hexactinellida are able to perform such internal contractions or not has not been proven yet. However, it seems to be unlikely. If Hexactinellida do not contract, the question arises, whether (1) contraction is only a character of all Metazoa other than Hexactinellida or if (2) contraction in Metazoa is a primary character and Hexactinellida lost it secondarily. The later is more likely, since cellular contractility is also present in unicellular eukaryotes. Movement of adult sponges on the substrate, in comparison, has only been demonstrated in demosponges (e.g. Bond 1992, Bond and Harris 1988, Fishelson 1981, Lieberkühn 1863, Nickel 2006, Pronzato 1999). It is likely that this will also be found in Homoscleromorpha, when investigated carefully under controlled conditions or in situ. Locomotion of adult or juvenile Hexactinellida or Silicispongia has not been reported to date. In contrast to contraction, which is quite widely distributed among sponges, locomotion in sponges seems to be an adaptation to unpredictable or unstable environments. This applies to many juvenile sponges, which have limited control over their environment chosen by phototactical or even chemotactical guidance of the larva prior to metamorphosis (e.g. compare Lieberkühn 1863 for movements of juvenile freshwater sponges). It also applies to moving species like Tethya spp. (Fishelson 1981, Sarà et al. 2001), presumably living in shallow coral reef communities or lagoons, where current, sediment and nutrient situation might fluctuate quickly on the microhabitat scale. Similar arguments can be addressed for light dependent species with associated photosynthetic microorganisms, like Chondrilla nucula SCHMIDT 1862 (Pronzato 2004), for which the light regime might change during the season, depending on growth competition with surrounding macroalgae. Biomechanics of contraction As shown above (see Sponge body contractions: Perception in modern biology), the question on the biomechanical mode of operation of contraction in sponges was frequently addressed and discussed contradictorily over the decades. However, most authors only focused on half of the biomechanical process, the contraction itself by the agonist system. Nevertheless, a further mechanism expanding the sponge during the second part of a contraction cycle has to be considered: the contraction antagonist. For the contraction agonist, three mechanisms are possible and discussed: (1) mesohyle contraction by actinocytes (e.g. Elliott 2004, Pavans de Ceccatty 1960); (2) epithelial contraction by endopinacocytes (e.g. Bagby 1970, Leys 2007, Leys and Meech 2006, Nickel et al. 2006c);

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

99

or 3. a combination of both (e.g. Pavans de Ceccatty 1981, 1986). Present experimental data and theoretical biomechanical implications suggests that a third mode is most realistic: (1) contractile filaments are present in pinacocytes (Bagby 1970, Leys and Meech 2006, Matsuno et al. 1988, Pavans de Ceccatty 1981, 1986); (2) the density of actinocytes in the mesohyle of highly contractile sponges is not high enough to explain alone the strong contractility, leading to volume reductions of up to 70% (Nickel 2004); (3) the contraction primarily leads to a reduction of the volume of the canal system, while the mesohyle does not or only minimally contracts (Nickel et al. 2006c, Nickel 2010). All this evidence suggests that pinacocytes are indeed the major contractile agonists. What about the antagonist system? If we regard the pinacocytes as the agonist, then we have to consider the following: a re-expansion of the pinacocytes after contraction would not necessarily produce the forces needed to reverse the narrowing of the aquiferous system elements. However, if one of the antagonist candidates is the system of choanocytes building up a water pressure inside the canals, then a second putative antagonist system is the mesohyle itself. Recent x-ray microtomography research demonstrated that during sponge contraction in T. wilhelma some regions of the mesohyle expanded rather than contracted (Nickel et al. 2006c, Nickel 2010). As a result, the network of extracellular matrix filaments is expanded, which was quantitatively confirmed by scanning electron microscopy (unpublished data). This phenomenon of elastic energy storage applies especially to very compact and collagen rich mesohyle regions of low cellular density. For mesohyle regions of lower collagen content but higher cellular density, the cells themselves may play active roles as antagonists. Expansion of the canals and consequently the whole sponge body might be supported by actinocytes within the mesohyle. During body contraction the actinocytes might be expanded or even stretched together with the mesohyle. During body expansion actinocytes might actively contract. However this is a speculative model supported by indirect evidence only. At present, in vivo microimaging of contracting actinocytes is missing. The only direct observation of contracting actinocytes demonstrated that they are involved in the active displacement of spicules during the formation of body filaments in T. wilhelma (Nickel and Brümmer 2004). In fact, the kinetics of a complete contraction-expansion cycle in T. wilhelma (Nickel 2004) and Ephydatia fluviatilis (Weissenfels 1989, 1990) show a bi-phasic tendency for the expansion. This might be explained by a double-antagonist system: during the first phase of expansion, initiated by the reversion of the pinacocyte contraction, elastic energy stored in the mesohyle would be released quite quickly, leading to a first expansion phase. The second, slower phase of expansion, leading to the restoration

© 2011 by Taylor and Francis Group, LLC

100

Key Transitions in Animal Evolution

of the full sponge body volume, would be caused by the pressure inside the aquiferous system produced by the choanocytes. Biomechanics of locomotion The biomechanics of sponge locomotion isn’t as documented or understood like contraction. During locomotion over the substrate sponges do not alter the area of attachment to a significant degree. The first movement model was suggested by Fishelson (1981). His model is based on the assumption of muscle like forces created by actinocytes within body filaments extending from the globular sponge body to the surrounding substrate. This hypothesis was falsified by other authors who could not confirm any contractile tension within the filaments (Bond 1992, Bond and Harris 1988, Nickel 2006, Nickel and Brümmer 2004). It can be assumed that the movements of all sponges follow the same mechanisms, regardless of the presence of body filaments or not. Bond and Harris (1988) were the first to suggest an amoeboid-like movement. The model is similar to the movement of fibroblast cells, which constantly reorganizes the cytoskeleton inside the cell during movement (Boal 2002). The model suggests that the movement of the sponge body is based on a constant reorganization of cells within the body (Bond 1992, Bond and Harris 1988). This mechanism is not to be confused with the growth of biomass by proliferation. Such a mechanism might play a role in body shape changes of some sponge species over very long time periods as observed in natural sponge communities (e.g. Pansini and Pronzato 1990). However, even in these cases, constant morphogenesis by cell movement plays an important role. It was also suggested that sponge movement is dependent on body contraction. Such peristaltic modes of movement can be frequently found among invertebrates (Trueman 1975). However, this is not the case in sponges, where contraction and locomotion on the substrate are clearly independent (Bond and Harris 1988, Nickel 2006). Open questions on contraction and locomotion At present we can only exclude some hypothesis on the cellular mechanisms involved in sponge contraction and locomotion. It is clear, that no kind of muscular system is involved in either. We have seen that actinocytes, which have been morphologically compared to smooth muscle cells (e.g. Bagby 1966, Fishelson 1981), do not play a major role in contraction and locomotion. The ongoing genome project on Amphimedon queenslandica HOOPER & VAN SOEST 2006 (aka Reniera sp. JGI-2005 WGS, searchable and downloadable

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

101

at http://trace.ensembl.org/ and http://www.ncbi.nlm.nih.gov/Traces) will help to unravel cellular properties. This has been demonstrated in evolutionary developmental biology of sponges (Adamska et al. 2007a, Adamska et al. 2007b, Larroux et al. 2006). Using an in silico approach as described by Gauthier and Degnan (2008), preliminary local blast searches of the A. queenslandica WGS trace files using conserved sequences of smooth muscle cell typical proteins, like titin and desmin did not recover similar gene sequences (unpublished data). Multidimensional life cell imaging methods (e.g. Hejnol and Schnabel 2006, Mori 1994) will be needed to assist in detailed analysis of the cellular contraction and movement dynamics during body contraction and locomotion. Intermediate filaments as well as receptor proteins and extracellular matrix elements involved in contraction in sponges are hardly ever investigated. Immunohistochemical methods using specific antibodies against these targets, eventually based on subsequent genomic analysis, will have to be forced (for an example see e.g. Wiens et al. 2001). Integrative systems in sponges Physiology of paracrine integration in sponges As mentioned above, paracrine signalling is present in protists and is therefore evolutionary ancient. It is important to note that the release of any chemical substance by any cell is at first a result of the cell’s present biochemical status and thus the cell’s statement about it. Such a statement is per se not directed. It enters the level of a ‘signal’ only, if a ‘receiver’ is present, e.g. a receptor system on another cell. This principle holds true for uni- and multicellular organisms. Obviously sponges react to environmental factors in a variety of ways, like all animals. All of these reactions involve intra- or intercellular signaling pathways, including second messenger systems, e.g. the temperature signaling cascade (Zocchi et al. 2003, Zocchi et al. 2001). However, such physiological pathways are not directly connected to coordination in the sense of nervous or nervous-like systems. Yet, such systems have been linked to discussions on the evolution of the nervous system (Jones 1962, Leys and Meech 2006). For the sake of clarity I will avoid mixing these topics. At present, the physiology of sponge signal integration is mostly defined by pharmacological studies, on which I will be focusing on here. Earlier studies were reviewed by Jones (1962) as well as Leys and Meech (2006). However, in an extensive study by Ellwanger and Nickel (2006), quantitative data was added, which was not included in any previous reviews.

© 2011 by Taylor and Francis Group, LLC

102

Key Transitions in Animal Evolution

Among transmitter substances, Acetylcholine (ACh) is well investigated. In fact, the first pharmacological studies on receptors are related to the muscarinic and nicotinic Ach receptor types (Dale 1914). While the nicotinic receptors are ligand-gated ion channels, the muscarinic types are metabotrophic systems. They play important roles in the CNS of all phyla (Walker et al. 1996, Walker and Holden-Dye 1991). For sponges, specific acetylcholine esterase (AChE) activity has been found in Sycon sp. (Lentz 1966), Hippospongia communis (LAMARCK 1814) (Thiney 1972) and also in T. wilhelma (Nickel 2001). However, no AChE activity could be demonstrated in earlier studies, though several species were investigated: Spongilla lacustris (LINNAEUS 1759) (Mitropolitanskaya 1941), Scypha sp. (Bullock and Nachmansohn 1942), Leuconia asperta (SCHMIDT 1862) and Siphonochalina crassa TOPSENT 1925 (Bacq 1947). On the other hand, in physiological studies by Pavans de Ceccatty (Pavans de Ceccatty 1971), ACh increased the rhythm and intensity of contraction in H. communis. AChE activity is also in T. wilhelma cells (Nickel 2001). In contrast to these results, physiological tests did not reveal a significant correlation between ACh and contraction. ACh did not induce contraction in T. wilhelma, but it did alter contraction frequency (Ellwanger and Nickel 2006). Eventually, it was shown that ACh is involved in the regulation of body extension formation and retraction (Nickel 2001). In the same study, nicotine did not affect contraction and rhythm in any way (Ellwanger and Nickel 2006). Even though the results on ACh are contradictory, it is likely that specific ACh esterases and receptors are present in at least Demospongiae and Calcispongia. Caffeine is a non-specific antagonist of adenosine-receptors in the mammalian brain. Its effect on the contractile behavior of T. wilhelma was also tested (Ellwanger and Nickel 2006). Caffeine induced contractions, increased the endogenous rhythm and attenuated the amplitude of endogenous contractions. Since the onset of caffeine-induced contraction seems to be decelerated, it must not act directly upon the contractile cells. From the disturbed endogenous rhythm and contraction amplitude, it seems likely that caffeine interferes with the regulation of endogenous contractions (Ellwanger and Nickel 2006). A putative adenosine receptor may be part of the pacemaker mechanism of endogenous contractions. In mammals, adenosine receptors are ubiquitous and important throughout all kind of tissues and cell types. They have also been shown to cause chronotropic effects in the heart (review in Klotz 2000). Adenosine receptors are all metabotrophic and link the ATP-driven energy metabolism to the signaling pathways of the cells. The adenosine signaling pathway strongly interacts with other messenger substances (Fredholm et al. 2000). Walker and co-authors (1996) concluded that such a central signaling mechanism evolved early and plays an important role in all invertebrates.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

103

Amino acid transmitters have also proved to be effective in sponges. In T. wilhelma, glycine induced effects are similar to caffeine: it stimulated contraction, fastened the endogenous rhythm, and attenuated the amplitude (Ellwanger and Nickel 2006). Together with GABA and glutamate, which are also effective in sponges (Ellwanger et al. 2006, Perovic et al. 1999, Ramoino et al. 2007), glycine is one of the more important amino acid transmitter in the vertebrate brain (Oberdisse et al. 1997). It acts as a transmitter in a wide range of invertebrates, e.g. in Hydra vulgaris PALLAS 1766 as an inhibitory transmitter (Pierobon et al. 2001). The results of Ellwanger and Nickel (2006) indicate that glycine is involved in the regulation of endogenous rhythm, not upon the contractile cells directly. Similar results can be found for GABA- and glutamate-induced contractions in T. wilhelma (Ellwanger et al. 2006). From these results, together with the characterization of a GABA/ glutamate-like receptor gene in Geodia cydonium (Perovic et al. 1999), the observation of a slower current under GABA exposure for Cliona celata (Emson 1966) and the immunohistochemical demonstration of GABAB receptors in Chondrilla nucula (Ramoino et al. 2007), it is evident that GABAergic signalling pathways exist in sponges. The catecholamine adrenaline is another substance, which was tested on sponges independently. A slight reduction of pumping rates was demonstrated in Cliona celata (Emson 1966). Lentz specifically stained adrenaline-containing bipolar and multipolar cell types in Sycon sp., using histochemical methods (Lentz 1966). In T. wilhelma, the strict limitation of effects to the elongation of endogenous contraction cycle duration in T. wilhelma suggests that adrenalin is directly involved or at least interfering with the regulation of the endogenous contraction rhythm (Ellwanger and Nickel 2006). Adrenaline is known to upregulate genes of the molecular clock in mammals and is therefore involved in the feedback loop regulation of circadian rhythm (Shibata 2004). In this context, it is interesting that both, Reiswig and Nickel, independently reported on diurnal rhythms in Cryptotethya sp. and T. wilhelma (Nickel 2004, Reiswig 1971). In conclusion, adrenaline might directly be involved in the regulation of biological rhythms in sponges. The biogenic amine serotonine was demostrated to be present in sponges using histochemical and imunohistochemical methods in Sycon sp. and Tedania ignis (Lentz 1966, Weyrer et al. 1999a). However, Emson found no effect upon Cliona celata (Emson 1966). In contrast, serotonine did immediately induce contraction in T. wilhelma (Ellwanger and Nickel 2006). The effect is limited: the endogenous contraction rhythm is not affected. However, the widespread findings of serotonine in all phyla of the metazoa, acting via ionotrophic and metabotrophic receptor types, points to an early evolution of this central signalling system (Walker et al. 1996, Walker and Holden-Dye 1991).

© 2011 by Taylor and Francis Group, LLC

104

Key Transitions in Animal Evolution

Nitric oxide (NO) is a gas which plays a central role in the regulation of a variety of physiological processes in vertebrates and invertebrates (Colasanti and Venturini 1998, Jacklet 1997). It interacts with a soluble guanylate cyclase (sGC) and is released by nitric oxide synthase (NOS) during conversion of l-arginine to l-citruline, mainly regulated by intracellular Ca2+. NO diffuses freely within the animal body and its range is limited by its short half life. The molecule is supposed to be an ancient auto- and paracrine messenger (Walker et al. 1996). NOS was shown to be present in the sponge Axinella polypoides SCHMIDT 1862, and it was involved in the temperature signalling cascade (Giovine et al. 2001). Ellwanger and Nickel demonstrated that NO induces contraction in T. wilhelma, but also modulates the endogenous contraction rhythm and amplitude. Due to its fast diffusion, the short half life and the limited range, NO is an optimal candidate for a contraction-inducing messenger, which acts in an auto- and paracrine manner. It is likely involved in feedback loops in the cellular pacemaker machinery of endogenous rhythm (Ellwanger and Nickel 2006). Contractions in sponges were also induced by cyclic AMP (Ellwanger and Nickel 2006). This is clearly an extracellular signaling event, which is most likely modulated by a specific cAMP receptor, similar to Dictyostelium discoideum RAPER 1935 (Lusche et al. 2005). Extracellular cAMP plays several regulative roles in Porifera: it is involved in the regulation of production and development of gemmules in freshwater sponges (Simpson and Rodan 1976), and it directly affects the locomotion of dissociated cells of Clathrina cerebrum (HAECKEL 1870) (Gaino and Magnino 1996). It may therefore link cellular mobility and body locomotion with contraction in sponges. All three phenomena are important aspects in the life of Tethya (Bond 1992, Bond and Harris 1988, Fishelson 1981, Nickel 2001, 2004, 2006). Summarizing this still rather patchy image of paracrine integration in sponges, it is safe to conclude that a variety of paracrine pathways are present in sponges, even though the behavioral repertoire of sponges is relatively low. However, transmitter substances are known to play important roles in unicellular eukaryotes, which tells us that these paracrine pathways are evolutionary ancient (Walker et al. 1996, Walker and Holden-Dye 1991). Most likely, they first acted as autocrine or intracrine messenger in unicellular eukaryotes (for the principal compare e.g. Ramoino et al. 2003, Ramoino et al. 2004, Ramoino et al. 2005). Consequently, it is not surprising to find such mechanisms in action in sponges. However, a lot of integrative research (i.e. physiological, molecular and morphological studies) will be necessary to unravel the details of these signaling networks.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

105

Physiology of electrical integration in sponges Apart from the extensive work on electrical integration in Hexactinellida, reports on this type of integration in other poriferans are rare. The reports of action potentials and impulse conduction in glass sponges are well documented and reviewed elsewhere (Leys and Mackie 1997, Leys and Mackie 1999, Leys et al. 1999, Leys et al. 2007, Leys and Meech 2006, Mackie et al. 1983). For this reason, I will not go into further detail on this subject. Although the demonstration of action potentials in hexactinellids is spectacular per se, it does not provide the key for understanding the early evolution of the nervous system. Clearly it demonstrates that voltagegated ion channels are present in Hexactinellids and most likely also in all other sponges. However, because the action potential travels through a syncytical tissue, signal transduction is not related to synaptic signal transition. This is displayed by the nervous system typical sequence of a double transition from an intracellular electric signal to an intercellular chemical signal and back to an intracellular electrical system again. On the other hand, action potentials and specific ion channels are known from protists, plants and bacteria (e.g. Mackie 1970, Schmidt-Rhaesa 2007, Schultz et al. 1986), so they are evolutionary ancient, too. For this reason, it is highly likely, that specific mechano-gated, voltage-gated and ligandgated ion channels are present in all sponges. No intracellular electrical recordings have been recorded so far from sponges except Hexactinellida. This is most likely due to problems related to cell size and the presence of mineral spicules in the tissue. However, experiments altering ion concentrations brought up evidence for the involvement of ion channels. Investigations upon the influence of extracellular ion concentrations pointed to the dependence of oscule contractions upon mono- and bivalent cations with interchangability for Na+, K+ and L+, respectively Ca2+, Mg2+ and Sr2+ in Microciona sp. (Prosser 1967, Prosser et al. 1962). In contrast, Na+ was not substitutable by K+ and Mg2+ or by Ca2+ in S. officinalis (Pavans de Ceccatty 1971). The direct application of electrical currents resulted in oscular contractions in the demosponge Ephydatia fluviatilis (McNair 1923). Changes in cell potential are therefore likely to play a role in signal integration in all sponges. For this reason, electrical synapses (gap junctions) might also be involved in signal transduction, at least temporarily. Macke (1970) suggested that electrical synapses (gap junctions) likely predated chemical synapses in evolution and may have been the foundation of a neuroid pre-nervous system. This view was supported by Loewenstein (1967) who demonstrated the electrical coupling of Haliclona sp. cells. Both works resulted in the intensified search for gap junctions in sponges.

© 2011 by Taylor and Francis Group, LLC

106

Key Transitions in Animal Evolution

Although some ultrastructural evidence was presented for gap junctions in sponges (Gaino and Sarà 1976, Green and Bergquist 1979) it was not confirmed by the same and other researchers (Garrone and Lethias 1990, Garrone et al. 1980, Green and Bergquist 1982, Lethias et al. 1983). The presence of gap junctions in sponges remains an open question. Morphology of integration systems in sponges The morphology of a putative integrative system in sponges is still a matter of speculation. As described above (see The quest for the elementary nervous system), the observation of contraction led to a number of histological studies after 1880 aimed at finding the contractile cells and nerve cells in sponges. Early evidence was presented for bipolar and multipolar cells interpreted as sensory and ganglion cells (Lendenfeld 1889), fusiform cells seen as nerve cells (‘aesthocytes’, Sollas 1888), flask-like cells regarded as sensory cells (Dendy 1891) and so on. It can be safely stated that at least Demospongiae and Calcispongia posses a number of cell types which morphologically resemble cell types of the nervous system in other metazoan groups (for a compendium see Boury-Esnault and Rützler 1997, De Vos et al. 1991, Harrison and De Vos 1991). In most cases morphology was the only evidence for attributing functions of an integrating system (for a detailed review see Jones 1962, Lentz 1968). Several histochemical methods which are used to specifically stain nerve cells (e.g. silver, Masson’s trichrome, etc.) were found to stain a variety of sponge cell types in Demospongiae (Pavans de Ceccatty 1955, 1959, 1960) and Calcispongia (Pavans de Ceccatty 1955, Tuzet et al. 1952). Consequently they were regarded as nerve-like cell by several authors, even if the morphology did not match any known cell types of the nervous system of other animal groups (Jones 1962). Some more specific evidence was presented by Lentz (1966, 1968), who used neurochemical methods to specifically demonstrate neurosecretory substances or enzymes in sponge cells. His results on acetylcholine esterase activity and the presence of serotonine in sponges was confirmed by others (Nickel 2001, Weyrer et al. 1999b). Although these results demonstrate that biochemical modules found in nervous systems are present in sponges, it does not necessarily mean that it is correct to regard these cells as nerve cells. More specific studies, combining immunohistochemistry and neurochemistry, are needed to reveal a more detailed image of how these modules are morphologically distributed among sponge cells. Molecular biology of integration in sponges A number of recent studies have focused on characterizing sponge genes, othologous or paralogous to nervous system specific genes of other

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

107

metazoa. One of the earliest examples is the molecular and functional characterization of a metabotrophic glutamate/GABA receptor type in Geodia cydonium (Perovic et al. 1999). The authors concluded that “these findings suggest that the earliest evolutionary metazoan phylum, the Porifera, possesses a sophisticated intercellular communication and signaling system, as seen in the neuronal network of higher Metazoa”. The concept of demonstrating typical molecular modules of the nervous system in sponges tempts to deduce a general architecture of an evolving nervous system in the ‘Urmetazoa’ or the LCAM. However, this is a path of trial and tribulation. As long as the detailed function of such sponge genes and their corresponding proteins within the cells and tissues are not characterized under a proper consideration of the morphological, ultrastructural and physiological context, the conclusions can only be preliminary. The same basic conditions apply for genome or large scale EST sequencing projects. An EST project on the homoscleromorph Oscarella carmella MURICY & PEARSE 2004 provided support for the presence of components of vesiclerelated signaling pathways (Jacobs et al. 2007, Nichols et al. 2006). These included synaptogamin, SNARE-complex components similar to syntaxin, neurocalcin, as well as a slit-gene, typically involved in axon guidance (Jacobs et al. 2007). The authors of these reports stated that “these recent observations in sponges suggest the high activity of equipment involved in vesicle transport, and the presence of some synaptic and developmental signaling components typically associated with bilaterian neural systems. Given that sponges lack formal synapses, it is worth noting that non-synaptic communication between cells via calcium waves can occur through a variety of mechanisms.” (Jacobs et al. 2007). As the authors correctly add, these genes also represent other functions in eukaryotic organisms. Final conclusions will have to wait until expression studies and pharmaceutical tests provide further supporting evidence. By analyzing the genomes of demosponge Amphimedon queenslandica and the cnidarian Nematostella vectensis STEPHENSON 1935, Sakarya et al. (2007) demonstrated the presence of 36 members of gene families involved in the post-synaptic scaffold in so called ‘lower metazoans’. Amongst them are voltage gated ion channels, ionotropic and metabotropic glutamate receptors. Obviously aware of the problem stated above, the authors cautiously stated that “Highly conserved protein interaction motifs and co-expression in sponges of multiple proteins whose homologs interact in eumetazoan synapses indicate that a complex protein scaffold was present at the origin of animals, perhaps predating nervous systems. A relatively small number of crucial innovations to this pre-existing structure may represent the founding changes that led to a post-synaptic element”. The first assumption that the presence of the molecular modules of the scaffold

© 2011 by Taylor and Francis Group, LLC

108

Key Transitions in Animal Evolution

predates the nervous system will have to be kept in mind. In fact, as we will see, this is a prerequisite for the evolution of nervous system sensu stricto. On the other hand, whether or not the presence of these proteins is equivalent to the presence of a highly organized structure like the post-synaptic scaffold is subject to discussion. I will return to this point in my consideration on the early evolution of a synapse-less pre-nervous integration system during phylogenesis of the Metazoa. Sakarya et al greatly demonstrated the value of genomic studies for reconstructing the biochemical repertoire of a theoretical LCAM in silico. This will inspire and provide a working hypothesis for a number of functional and morphological studies to further characterize the obvious integrative capacity of pre-nervous systems in those metazoan groups which first branched off during evolution. Co-Evolution of Contraction and Integration: Pre-Nervous Milestones in Sponges The lack of early evolutionary scenarios The evolution of the nervous system has been discussed extensively in the last century by numerous authors from Parker (1910) to Meech and Mackie (2007). Nevertheless, there is still a significant lack of theories on the early evolution of integrative systems in Metazoa. Most of the publications focus on the evolution of excitability in Metazoa. As a result, the neuron concept (i.e. Barbara 2006, Cajal 1937) has played a major role in hypotheses on the nervous system evolution. This observation is valid for all major reviews and textbooks on neurobiology (e.g. Bullock and Horridge 1969, Grundfest 1959, 1965, Hanström 1928, Jones 1962, Leys 2007, Leys and Meech 2006, Mackie 1979, 1990, Meech and Mackie 2007, Pantin 1952, 1956, Parker 1910, 1919, Passano 1963, Schmidt-Rhaesa 2007). Consequently, most authors focus on the structural evolution of complexity in the nervous systems of bilaterian animals, trying to reconstruct “the nervous system of Urbilateria” (Ghysen 2003). The reason for this situation is probably best understood by referring to Bullock’s and Horridge’s (1969) statement in their attempt to present ‘defining features of a nervous system’: “A nervous system may be obvious and easily identified, as in higher groups, at least in its central parts; or it may be extremely difficult either to recognize or to deny, as in the multicellular sponges and the highly coordinated ciliate protozoans (heterotrichs).” By referring to the concept of ‘dependant effectors’ of Parker (1910, 1919), Pantin (1952, 1956) and Passano (1963), they continue to argue that “The relevance of the definition for sponges, and other metazoans in which a nervous system is to be demonstrated, is that it provides anatomical and physiological criteria to be satisfied by any true nervous system. Thus presumptive nerve cells must be shown to

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

109

connect receptor sites with effectors or to connect with each other. Cells that send prolongations to end in open tissue or spaces or on spicules are unlikely to be nervous, unless other argument clearly exist.” (Bullock and Horridge 1969). Even though it is not clearly stated by most authors, they all assume the presence of chemical synapses in combination with electrical conductions over long distances. This should be a major defining character of any ‘true nervous system’ (i.e. Bullock and Horridge 1969, Grundfest 1959, Hanström 1928, Schmidt-Rhaesa 2007). This is influenced by the ‘neurone doctrine’ of Ramon y Cajal and Heinrich Wilhelm von Waldeyer-Hartz (for historic overviews see i.e. Barbara 2006, Smith 1996). A historic overview on basic concepts of nervous system evolution is given by Hanström (1928). Basically, all of the theories center on the differentiation of ‘receptor’ and ‘effector’ elements in the first nervous systems. Oscar and Richard Hertwig (1878) assumed that muscle cells, sensory cells and ganglion cells evolved from indifferent epithelial cells under constant interactions. Parker (1919) suggested that independent primitive receptor-effector cells received stimuli and react by appropriate activity, i.e. contraction. Parker assumed that in the course of metazoan evolution, receptors and effectors were separated in space and structure, but remained connected through axons. Although the term ‘synapse’ and the connected concept was given earlier (Sherrington 1897), it did not play a major role in the Parker’s and subsequent theories. But which evolutionary sequence resulted in this morphologically and physiologically highly specialized system? Grundfest (1959) suggested that neurosecretion—and not contraction—represents the ancient effector activity of Parker’s primitive receptor-effector cells. He continues that “Each cell [of receptor and effector type] retained its activity both as a receptor and effector, though maintaining these functions in a modified and often specialized form.” (Grundfest 1959). Lentz referred to Grundfest’s conclusions when he reported on transmitter activity in sponges (Lentz 1966, 1968). However, from his results he concluded that synapses must be present in sponges and would have to be confirmed ultrastructurally or morphologically. At present, there is no ultrastructural evidence for synapses in sponges (Jones 1962, Leys et al. 2007, Mackie 1990, Meech and Mackie 2007, Simpson 1984). Due to genomic research (see above: Molecular Biology of Integration in Sponges) and the finding of a high number of genes related to the post-synaptic scaffold (Sakarya et al. 2007) and other synapse-related genes (Jacobs et al. 2007), the idea prevails that sponges might possess synapse-like cellular junctions: “The core potential for evolving synapses in sponges may extend to other types of junctions.” (Sakarya et al. 2007). Statements like this reveal quite a traditional view on the evolution of the nervous system, which seems

© 2011 by Taylor and Francis Group, LLC

110 Key Transitions in Animal Evolution not to plumb the depths of possible scenarios. In this case, it is implied that synapses evolved from other type of cell junctions. Such a scenario is unlikely, given the ultra- and nanostructural and thus genetic differences between synapses and cell junctions like gap junctions, tight junctions, or desmosomes. In the following paragraphs, I will present a hypothesis on the early evolution of a (pre-)nervous integration system combining the views of Grundfest (1959), with those on compartmentalization of signaling in neurons (Mattson and Bruce-Keller 1999), genomic evidence (Jacobs et al. 2007, Sakarya et al. 2007), histological and morphological evidence (as reviewed by Jones 1962, Pavans de Ceccatty 1979, 1989, Simpson 1984) and physiological evidence (for details see Ellwanger et al. 2006, Ellwanger and Nickel 2006, Lentz 1966, 1968, Leys and Meech 2006 and other authors referenced in here). The Paracrine-to-electrochemical-dominance transition hypothesis Axioms The following conclusions from the literature discussed above will serve as axioms for the hypothesis on the early evolution of the nervous system in Metazoa presented in the following two subchapters: 1. Unicellular eukaryotes represent independent effectors or receptor-effector cells sensu Parker (1919). Explanation: ciliates are capable of producing action potentials by way of voltage-gated ion channels as an answer to external stimuli (Schultz et al. 1986). 2. Unicellular eukaryotes use auto- and paracrine signaling pathways for intercellular communication. Explanation: The release of signalling substances as statements of the biochemical status of a cell is pleisiomorphic. Ciliates and other protists are able to release several signaling substances and posses receptor systems for specific transmitter substances (Ramoino et al. 2003, Ramoino et al. 2004, Ramoino et al. 2005, Walker et al. 1996, Walker and Holden-Dye 1991). 3. The sponge body constitutes a network of dependent effectors or receptoreffector cells sensu Grundfest (1959). Explanation: Sponge cells are capable of producing and releasing neurotransmitters (Lentz 1966, 1968, Nickel 2001, Thiney 1972, Weyrer et al. 1999b). They possess the gene and protein repertoire for neurotransmitter receptor systems (Jacobs et al. 2007, Perovic et al. 1999, Ramoino et al. 2007, Sakarya et al. 2007). They specifically react upon neurotransmitters and agonist/antagonists of receptor systems involved in the nervous system (Ellwanger et al. 2006, Ellwanger and Nickel 2006, Emson 1966, Perovic et al. 1999). This, however, does not imply the presence

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

4.

5.

6.

7.

111

of synapses or direct cell contacts. The network is mainly connected by chemical messengers (Ellwanger and Nickel 2006). Contraction in sponges is mainly mediated by epithelial cells. Explanation: Pinacocytes are the main contractile effectors in sponges (e.g. Bagby 1970, Leys 2007, Leys and Meech 2006, Matsuno et al. 1988, Nickel et al. 2006c, Nickel 2010), eventually assisted by mesenchymal actinocytes (e.g. Pavans de Ceccatty 1981, 1986). No specialized muscles are present. Electrochemical potential changes coupled to intra and extracellular signaling pathways are a common character of all Metazoa. Explanation: Hexactinellids display action potentials (Leys 2007, Leys and Mackie 1997, Leys and Mackie 1999, Leys et al. 1999, Mackie et al. 1983), Demospongiae display potential changes (Carpaneto et al. 2003, Loewenstein 1967). Similar mechanisms are present in ciliates (Schultz et al. 1986). Trophic functions of neurotransmitter-specific receptor systems predate integrative functions in metazoan (pre-)nervous systems. Explanation: Neurotransmitters like GABA and glutamate are evolutionary ancient (Walker et al. 1996, Walker and Holden-Dye 1991). In Metazoa they are also involved in axon guidance (Ruediger and Bolz 2007) and trophic functions in the nervous system ontogeny (Nguyen et al. 2001), which are most likely evolutionary ancient functions, too. Synapses sensu stricto are an autapomorphy of the Eumetazoa. Explanation: Synapse structures are present in Cnidaria and Ctenophora as well as in all Bilateria (Bullock and Horridge 1969, Hanström 1928, SchmidtRhaesa 2007).

The hypothetical paracrine pre-nervous system in the first metazoa (LCAM) Unicellular eukaryotes like ciliates communicate via secretory pathways, coupled to metabotrophic intracellular signaling pathways and voltage-gated ion channels (Fig. 4A). Consequently, it is likely that the intercellular communication within the first Metazoa was based on the same mechanisms. Thus, a pre-nervous integration system consisting of receptor-effector cells sensu Grundfest (1959) must have been present from the beginning of multicellularity in the Metazoa. It can be regarded as a key feature of the Last Common Ancestor of Metazoa (LCAM). Its function must have closely resembled the auto- and paracrine signalling mechanisms used by unicellular eukaryotes, except that chemical messengers are released into the extracellular matrix and putative body cavities instead into the environment (Fig. 4B). Such extracellular but intracorporeal chemical signals dominate the whole integration system on the spatial

© 2011 by Taylor and Francis Group, LLC

112 Key Transitions in Animal Evolution

Fig. 4. Explanatory scheme presenting evolutionary levels as integral parts of the ‘paracrineto-electrochemical-dominance transition’ hypothesis. For details refer to the text. Diffusion gradients A. Level of auto- and paracrine signalling in unicellular eukaryotes. (1) External stimulus followed by (2) a intracellular signal leading to (3) the release of a chemical messenger mediating (4) autocrine and (5) paracrine reception. B. Hypothetical level of the LCAM: (1) a sensed extracorporeal stimulus results in (2) the release of a paracrine messenger into the extracellular matrix (blue gradient). This diffusing signal reaches receptive cells which act as receptor-effectors sensu Grundfest including (3) secretion of secondary chemical signals (integration) or the same chemical substance (positive feedback). These secondary signal gradients (orange gradients) differ from the primary signal gradient and eventually reach other cells. C. Hypothetical intermediate level, eventually present in Demospongiae: again (1) an external stimulus is received by epithelial cells, followed by (2) the release of a chemical messenger (blue gradient). This signal is received by neighboring epithelial cells and mesenchymal multipolar cells. It locally triggers the opening of ligand-gated ion channel in the receptor cell, which (3) results in a potential change within the whole cell or a spreading local potential change over the cell body (action potential). The incurrent of specific ions (e.g. Ca2+) and/or secondary messenger systems locally trigger the secondary release of extracellular messengers (orange gradients). During the proposed evolutionary transition, the secretory complexes are compartmentalized within the cells. The secondary signal is received by other cells and triggers secondary effects like (4) secretion, cell locomotion and (5) contraction of epithelial cells. D. Level of a simple nervous system representing the simple reflex-arc sensu Parker, from (1) the incoming external signal at epithelial receptor cells over (2) synapses to a nerve cell, generating excitatory or inhibitory postsynaptic potentials. This neuron (3) might integrate signals from separate sensory cells and generate an outgoing signal at (4) a synapse towards a muscle cell, representing a highly differentiated contractile effector cell (5). Color image of this figure appears in the color plate section at the end of the book.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

113

and temporal scale: signal molecules travel the distances between the cells by way of diffusion which is relatively slow. Consequently, chemical gradients of messenger molecules are present throughout the body. Morphology strongly influences the spatial structure of the signal’s diffusion gradients (e.g. cellular density, local extracellular matrix composition and density, body cavities, etc.). In this way the 3-dimensional diffusion gradients are shaped secondarily. Depending on differentiation, cells react upon incoming chemical signals by either the release of the same substances (positive feedback) or the release of other signal substances (chemical integration) including specific antagonists (negative feedback). More complex intracellular signaling pathways like molecular biological clocks (Ishida et al. 1999) lead to delayed release of substances (temporal integration). Cells dominated by such pathways can be regarded as the first pacemakers of the LCAM’s pre-nervous system. Electrochemical changes of the cell potential mediated by Ca2+- and presumably other ion channels are likely to have played important roles in the control of intracellular biochemical pathways (Leys and Meech 2006). If syncytical tissue was present in the LCAM to a larger degree—for which we have no evidence—electrical conduction might have played an early role in intrasyncytical communication over longer distances, like in Hexactinellida (Leys et al. 2007). However, this seems to be a derived character of the later. As stated by Grundfest (1959), contraction of certain specialized cells was the second effector mechanism in evolutionary early coordination systems besides the general ability of neurosecretion manifested in every cell type. In all probability, contraction was not the exclusive function of these cells. Instead, most likely the first contractile cells were the epithelial cells of the LCAM. The cellular contractile apparatus did not resemble the highly ordered, fast acting contractile intermediate filament system of muscle cells, but were less specialized like contractile endothelial cells of vertebrates or pinacocytes of sponges. Recent results on morphology and physiology of integration systems in demosponges suggest that such a non-directional paracrine pre-nervous system is the main integration system (Ellwanger and Nickel 2006). Whether or not some groups of Demospongiae sensu stricto, Homoscleromorpha or Calcispongia took further evolutionary steps as described below remains an open question to date. The paracrine-to-electrochemical-dominance transition At some phases during the evolution of the nervous system sensu stricto, (Bullock and Horridge 1969, Hanström 1928) selective pressures must have favored (1.) directed communication and (2.) faster communication

© 2011 by Taylor and Francis Group, LLC

114 Key Transitions in Animal Evolution (in comparison to diffusion of molecules). This lead to further cellular specialization and compartmentalization as well as a transition of the temporal and spatial dominance from paracrine signal transduction to electrochemical signal transduction. Contrary to Parker (1919), and following Hertwig and Hertwig (1878), I assume a parallel evolution of highly specialized contractile effectors (muscle cells), sensory cells and ganglion cells from non-specialized epithelial cells. A variety of selective pressures must have favored compartmentalization of the body and faster contractile responses to external stimuli. Namely, the spatial concentration of contractile cells into defined cellular layers and eventually contractile cellular sheets must have privileged a co-evolution of faster contraction mechanisms and directed intercellular communication. It is likely that the evolution of these cellular differentiations took place parallel to the evolution of the mesoderm. However, this will have to be proven, since the evolutionary origin of the mesoderm is presently under discussion (Martindale et al. 2004, Seipel and Schmid 2006, Spring et al. 2002). The trophic and morphogenetic effects of several neurosecretory molecules predated the evolution of their transmitter function. They modulated growth and elongation of cells, frequently guided by gradients of neurosecretions. Like all other cells, these elongated cells were capable of reception and neurosecretion by itself. If faster signaling is favored by evolutionary pressures, slight accidental structural accumulation of membrane bound and cytoplasmic signaling pathway proteins would have enhanced the speed of intracellular signal transmission. Such intracellular accumulation might have been influenced by the mentioned neurosecretory cell guidance effects. However, slight unequal distributions of secretion and receptor complexes would lead to an alteration of the neurosecretion gradient fields around these cells (Fig 4C). The elongation of cells and the concentration of the neurosecretory machinery towards the cellular ends in conjunction with electric potential changes within the cells would have resulted in several benefits: (1) cost-benefit for neurosecretion would increase, since it is localized; (2) the temporal dominance of messenger diffusion would be balanced by a higher spatial share of the faster intracellular electric conduction; (3) autocrine effects (external feedback loops) would be minimized. A decrease of distance between the site of neurosecretion and putative receptor sites in neighboring cells would result in further effects: (1) it would increase signal specificity; (2) it would further increase cost benefit; due to a decreasing dilution over diffusion distance, it would be possible to obtain the same or even higher levels at the receptor sites, by releasing lower messenger concentrations; (3) autocrine effects would be completely eliminated, enabling strictly directed signal transduction between single cells.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

115

Gradually such an evolutionary scenario would favor further compartmentalization within the cell and thus slowly result in the formation of pre-synaptic structures and finally synapses (Fig. 4D). During this evolutionary course, the dominance of paracrine signal transduction by means of time and distance is broken. In other words, the temporal and the spatial share of the paracrine subsystem of an evolutionary early (pre-) nervous system is minimized. Instead, the faster intracellular electrical signal transduction covers larger distances between the cells thus gaining temporal and spatial dominance within the metazoan integration system. This scenario is a prerequisite for the evolution of spatial focused two-waytransmission as found in the nerve nets of cnidarians and ctenophorans. This is also true for the polarized one-way-transmission which dominates centralized nervous systems (Anderson 1985, Bullock and Horridge 1969, Meech 2008). Phylogenetic Considerations and Open Questions This leads me to the obvious question: can we provide phylogenetic evidence for this scenario? Are we able to identify groups among recent Metazoa which can be regarded as representatives of intermediate stages in the paracrine-to-electrochemical-dominance transition? The fact that sponges possess synaptic protein complexes but do not display functional synapses qualifies them as candidates. Clearly, all higher Taxa —Hexactinellida, Demospongiae sensu stricto, Homoscleromorpha and Calcispongia—display pre-nervous integrative systems. However, we still know less about these systems in comparison to other integrative systems, like the nerve-nets of cnidarians. We will need to include expression studies as well as immunohistochemical cLSM and TEM studies to demonstrate the distribution of synaptic proteins among sponge cell types, and especially on the sub-cellular level. The phylogenetic relations among all of the basal branching metazoan groups will also be essential. The phylogenetic position of Placozoa will be of special interest (e.g. Collins et al. 2005, Schierwater and DeSalle 2007), as well as the scenario of paraphyly among sponges (e.g. Borchiellini et al. 2001). Neither groups display nervous systems (Fig. 5a). At present, the literature on integrative systems in sponges hardly distinguishes any results between Hexactinellida, Demospongiae, Homoscleromorpha and Calcispongia. This will have to be sorted out. The paraphyly scenario that has each of these four clades branching out separately from the metazoan stem lineage, as well as the possibility of a sponge-like LCAM implies far-reaching possibilities. Each of the four taxa might in fact represent different graded stages of the early evolution of the pre-nervous system. Studies involving members of all taxa are necessary to complete this

© 2011 by Taylor and Francis Group, LLC

116 Key Transitions in Animal Evolution picture. On the other hand, recent phylogenomic anlyses supported the classic view of monophyletic Porifera (Philipp et al. 2009, Pick et al. 2010). In this case it will be interesting to investigate which of the poriferan prenervous system modules are dapomorphies and which arr plesiomorphies inherited from the LCAM (Fig. 5b).

Fig. 5. Summary of the co-evolution of integration and effector systems in Metazoa. The phylogenetic tree follows (a) Erpenbeck and Wörheide (2007), Borchiellini et al. (2001) and others and (b) Philippe et al. (2009) and Pick et al. (2010). The evolution of the ultrastructure of synapses can presently not be resolved, but the synapse gene repertoire must be pleisiomorphic to all Metazoa. In this context, the relative phylogenetic position between sponges and the Placozoa is not crucial and resolving this problem will most likely not add any information to the question of synapse evolution. In contrast, the phylogenetic position of the Ctenophora provides essential information, since they display true synapses. For further details refer to the text.

Recently, Dunn et al. (Dunn et al. 2008) suggested that Ctenophora might represent the most basal branching metazoan clade. The support for this hypothesis is relatively weak at the moment. However, it is worth thinking about it in the context of nervous system evolution. Since ctenophorans posses synapses and nerve cells are arranged in a nerve nets (Bullock and Horridge 1969, Hernandez-Nicaise 1973, Schmidt-Rhaesa 2007) the result would be that it either evolved independently or that the LCAM possessed a synaptic nervous system. In this case, the synapseand nerve-less condition is the sponge clades and the Placozoa would be secondary. This scenario seems to be less parsimonious than others, which consider sponges and Placozoa to represent the basal branches of the metazoan tree (Fig. 5).

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

117

Conclusions and Outlook The history of perception of sponges as animals is full of controversial discussions and miss-interpretations. Interestingly, contraction and sensibility, or in other words, the questions of reception, integration and effector systems, have all played a major role in this controversy. As soon as the fine structure of nerve cells was described, the controversy became one on the presence of nerve cells in sponges. The statement of Jones (1962) that there is no nervous system in sponges subsided the discussions. The question of the presence of absence of a nervous system in sponges still arises regularly. Taking into account the behavioral, physiological, morphological, histological and genomic data on sponges reviewed above, my personal impression is that the question itself is the wrong one: “Is there a nervous system in sponges?”—This can clearly be denied based on present data. But what is the informative value of this answer?—Almost none. Understanding the evolution of biological systems does not work simply on the basis of present vs. absent statements. If the nervous system evolved, and it did, we will be able to find traces of the early evolution. Consequently, the question(s) will have to be different and probably more specific: Which nervous-system typical modules or pre-cursors of such modules can be found in those animal groups which do not possess a nervous system in the strictest sense? We are talking about all levels from genomics to the tissue or animal level. These questions have guided me when considering a pre-nervous integration system in sponges (Ellwanger and Nickel 2006 and this publication). Though this pre-nervous system is at present not yet well enough characterized in detail, it is possible to outline it on an integrative data basis. Clearly, recent genomic research (Jacobs et al. 2007, Nichols et al. 2006, Sakarya et al. 2007) has greatly stimulated the genesis of the ‘paracrine-to-electrochemical-dominance transition’ hypothesis of the early evolution of the nervous system from the pre-nervous system. The open questions regarding the hypothesis presented here or other hypotheses on the early evolution of the nervous system will not be solved by pure genomic or other purely molecular oriented approaches (Jacobs et al. 2007, Sakarya et al. 2007), nor by pure physiological approaches (e.g. Elliott and Leys 2007, Ellwanger et al. 2006, Ellwanger and Nickel 2006). Ideally, future research will have to combine both approaches and add a third component: morphology. In situ hybridization as well as life cell imaging techniques in combination with physiological and (cell/tissue) behavioral studies will play major roles in an integrated approach. To give an example: The presence of certain genes related to the nervous system (Sakarya et al. 2007) or even the expression of some receptors (Perovic et al. 1999) in sponges do not necessarily exhibit the same functions like

© 2011 by Taylor and Francis Group, LLC

118 Key Transitions in Animal Evolution those in the nervous system of other metazoans. Regardless of whether or not the target genes are orthologs or paralogs of typical nervous systemrelated genes of other metazoans, these genes might represent different functionalities. Many elements of the nervous system are known from unicellular eukaryotes, e.g. the GABA-receptors in Paramecium caudatum MÜLLER 1773 (Ramoino et al. 2003, Ramoino et al. 2004, Ramoino et al. 2005). Consequently, we can expect per se to find similar genes in sponges. Thus it will be most important to support detailed genetic, functional and morphological characterizations of the contraction effectors, pacemakers of endogenous rhythms, and the signaling pathways in sponges. For this, phylogenetic considerations have to be taken into account. Ideally, such studies would include model species ranging from demosponges (e.g. T. wilhelma, E. fluviatilis, S. lacustris and others) to homoscleromorphs (e.g. Oscarella spp.) and calcareous sponges (Clathrina spp., Sycon spp. and others). Only in this way, can we expect to gather sequential evolutionary information on the early metazoan (pre-)nervous signalling system and the associated (pre-)muscular contraction systems. Finally, it is interesting to point out that the pre-nervous system in sponges is strongly associated with the sponge epithelia, the pinacoderm. It was previously suggested that the early evolution of central nervous system brought about an “era of skin brains” (Holland 2003). By referring to the epithelia of Hydra Meinhardt (2002) stated that “an old body became a young brain”. Following this perspective, we might regard the pinacoderm of sponges as an evolutionary ‘very young skin brain’. Acknowledgements I thank Sally P. Leys (Edmonton, Canada), Dennis V. Lavrov (Ames, USA), Calhoun Bond (Greensboro, USA), Roberto Pronzato (Genova, Italy), Werner E.G. Müller (Mainz, Germany) as well as George O. Mackie and Henry Reiswig (both Victoria, Canada) for inspiring this work and for discussions and comments on the topic. I am grateful to Martin S. Fischer (Jena, Germany), Hans-Dieter Görtz and Franz Brümmer (both Stuttgart, Germany) for supporting my work and providing infrastructure. I thank Bernd Schierwater (Hannover, Germany) and Rob de Salle (New York, USA) for organising the SICB-symposium, Angie Faust (Hannover, Germany) for assistance during the editorial process of this chapter, and Katherine Zocholl (Ulm, Germany) for English language editing. References Adamska, M. and S.M. Degnan, K.M. Green, M. Adamski, A. Craigie, C. Larroux, and B.M. Degnan. 2007a. Wnt and TGF-ß expression in the sponge Amphimedon queenslandica and the origin of metazoan embryonic patterning. PLoS ONE 2: e1031.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

119

Adamska, M. and D.Q. Matus, M. Adamski, K. Green, D.S. Rokhsar, M.Q. Martindale, and B.M. Degnan. 2007b. The evolutionary origin of hedgehog proteins. Curr. Biol. 17: R836–R837. Aelianus, C. De natura animalium libri septemdecim. F. Jacobs. [ed.] 1832. Frommann, Jena. Anderson, P.A. 1985. Physiology of a bidirectional, excitatory, chemical synapse. J. Neurophysiol. 53: 821–835. Anderson, P.A.V. [ed.]. 1990. Evolution of the First Nervous Systems. Plenum, New York. Ankel, W. 1948. Über Fragen der Formbildung und der Zell-Determination bei Süßwasserschwämmen. Verh. Dtsch. Zool. Ges. 1948: 58–66. Aristotle. De natura animalium libri novem; de partibus animalium libri quatuor; de generatione animalium libri quinqui. In: T. Gaza [ed.] 1498. Domini Octaviani Scoti, 89 pp. Aristotle. 1910. The History of Animals. Transl. by D‘Arcy Wentworth Thompson. Oxford: Clarendon Press. [Electronic reprint]. The University of Adelaide Library. http://etext. library.adelaide.edu.au/a/aristotle/history/ Arndt, W. 1941. Lebendbeobachtungen an Kiesel- und Hornschwämmen des Berliner Aquariums. Zool. Garten (Leipzig) 13: 140–166. Bacq, Z.M. 1947. L’acétylcholine et l’adrénaline chez lez Invertébrés. Biol. Rev. 22: 73–91. Bagby, R.M. 1966. The fine structure of myocytes in the sponges Microciona prolifera (Ellis and Sollander) and Tedania ignis (Duchassaing and Michelotti). J. Morphol. 118: 167–182. Bagby, R.M. 1970. The fine structure of pinacocytes in the marine sponge Microciona prolifera. Z. Zellforsch. Mikroskop. Anat. 105: 579–594. Barbara, J.-G. 2006. The physiological construction of the neurone concept (1891–1952). Comptes Rendus Biologies 329: 437–449. Barthélemy-Saint Hilaire, J. 1883. Histoire des Animaux D’Aristote [Electronic Ressource]. Libraire Hachette. http://remacle.org/bloodwolf/philosophes/Aristote/tableanimaux. htm. Bidder, G.P. 1937. The perfection of sponges. Proc. Linn. Soc. London 149th session: 119– 147. Boal, D. 2002. Mechanics of the Cell. Cambridge University Press, Cambridge. Bond, C. and A.K. Harris. 1988. Locomotion of sponges and its physical mechanism. J. Exp. Zool. 246: 271–284. Bond, C. 1992. Continuous cell movements rearrange anatomical structures in intact sponges. J. Exp. Zool. 263: 284–302. Borchiellini, C. and M. Manuel, E. Alivon, N. Boury-Esnault, J. Vacelet, and P.Y. Le. 2001. Sponge paraphyly and the origin of Metazoa. J. Evol. Biol. 14: 171–179. Borchiellini, C. and C. Chombard, M. Manuel, E. Alivon, J. Vacelet, and N. Boury-Esnault. 2004. Molecular phylogeny of Demospongiae: implications for classification and scenarios of character evolution. Mol. Phylogenet. Evol. 32: 823–837. Boury-Esnault, N. and K. Rützler. 1997. Thesaurus of sponge morphology. Smithsonian Contributions to Zoology 596: 1–55. Boury-Esnault, N. 2006. Systematics and evolution of Demospongiae. Can. J. Zool. 84: 205–224. Bowerbank, J.S. 1856. On the vital powers of the Spongiadae. Rep. Brit. Assoc. London 26: 438–451. Brøndsted, H.V. 1942. Formbildungsprozesse bei einem sehr primitiven Metazoon, dem Süsswasserschwamm Spongilla lacustris (L.). Protoplasma 37: 244–257. Bullock, T.H. and G. Horridge. 1969. Structure and Function of the Nervous Systems of Invertebrates. W.H. Freeman, San Fransico. Bullock, T.H. and D. Nachmansohn. 1942. Coline esterase in primitive nervous systems. J. Cell. Comp. Physiol. 20: 239–242. Burton, M. 1948. Observations on littoral sponges, including the supposed swarming of larvae, movement and coalescence in mature individuals, longevity and death. Proc. Zool. Soc. London 118: 893–915.

© 2011 by Taylor and Francis Group, LLC

120

Key Transitions in Animal Evolution

Carpaneto, A. and R. Magrassi, E. Zocchi, C. Cerrano, and C. Usai. 2003. Patch-clamp recordings in isolated sponge cells (Axinella polypoides). J. Biochem. Biophys. Methods 55: 179–189. Carter, H.J. 1848. Notes on the species, structure, and animality of the freshwater sponges in the tanks of Bombay (genus Spongilla). Annals and Magazine of Natural History 2nd. ser. 1: 303–311. Colasanti, M. and G. Venturini. 1998. Nitric oxide in invertebrates. Mol. Neurobiol. 17: 157– 174. Collins, A.G. and P. Cartwright, C.S. McFadden, and B. Schierwater. 2005. Phylogenetic context and basal metazoan model systems. Integr. Comp. Biol. 45: 585–594. Dale, H.H. 1914. The action of certain esters and ethers of choline and their relation to muscarine. J. Pharmacol. Exp. Ther. 6: 147–190. De Vos, L. and G. Van De Vyver. 1981. Étude de la contraction spontanée chez ‘éponge d’eau douce Ephydatia fluviatilis cultivée in vitro. Ann. Soc. Roy. Zool. Bel. 111: 21–32. De Vos, L. and K. Rützler, N. Boury-Esnault, C. Donaday, and J. Vacelet. 1991. Atlas of Sponge Morphology. Smithsonian Institution Press, Washington. Dendy, A. 1891. Memoirs: Studies on the Comparative Anatomy of Sponges: III.—On the Anatomy of Grantia labyrinthica, Carter, and the so-called Family Teichonida. Q. J. Microsc. Sci. 32: 1–39. Dunn, C.W. and A. Hejnol, D.Q. Matus, K. Pang, W.E. Browne, S.A. Smith, E. Seaver, G.W. Rouse, M. Obst, G.D. Edgecombe, M.V. Sorensen, S.H.D. Haddock, A. Schmidt-Rhaesa, A. Okusu, R.M. Kristensen, W.C. Wheeler, M.Q. Martindale, and G. Giribet. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–749. Elliott, G. 2004. The contractile apparatus of the sneezing freshwater sponge Ephydatia muelleri. Newsletter of the American Microscopical Society, Fall issue: 5–7. Elliott, G.R.D. and S.P. Leys. 2007. Coordinated contractions effectively expel water from the aquiferous system of a freshwater sponge. J. Exp. Biol. 210: 3736–3748. Ellis, J. 1755. An essay towards a natural history of the corallines and other marine products of the like kind, commonly found on the coasts of Great Britain and Ireland, London. Ellis, J. and D. Solander. 1786. The Natural History of Many Curious and Uncommon Zoophytes, Collected from Various Parts of the Globe. Systematically Arranged and Described by the Late Daniel Solander. Benjamin White & Son, London , 206 pp and 63 Pl. Ellwanger, K. and A. Eich, and M. Nickel. 2006. GABA and glutamate specifically induce contractions in the sponge Tethya wilhelma. J. Comp. Physiol. A 193: 1–11. Ellwanger, K. and M. Nickel. 2006. Neuroactive substances specifically modulate rhythmic body contractions in the nerveless metazoon Tethya wilhelma (Demospongiae, Porifera). Frontiers in Zoology 3: 7. Emson, R.H. 1966. The reactions of the sponge Cliona celata to applied stimuli. Comp. Biochem. Physiol. 18: 805–827. Erpenbeck, D. and G. Wörheide. 2007. On the molecular phylogeny of sponges (Porifera). Zootaxa 1668: 107–126. Esper, E.J. 1794–1805. Die Pflanzenthiere: in Abbildungen nach der Natur mit Farben erleuchtet nebst Beschreibungen. Raspe, Nürnberg. Fishelson, L. 1981. Observations on the moving colonies of the genus Tethya (Demospongia, Porifera): 1. Behavior and cytology. Zoomorphology 98: 89–100. Fredholm, B.B. and G. Arslan, L. Halldner, B.r. Kull, G. Schulte, and W. Wasserman. 2000. Structure and function of adenosine receptors and their genes. Naunyn Schmiedebergs Arch. Pharmacol. 362: 364–374. Gaino, E. and M. Sarà. 1976. Ultrastructural observation on the cell junctions in aggregates of Reniera fulva (Porifera, Demospongiae). Boll. Mus. Ist. Biol. Univ. Genova 44: 5–10. Gaino, E. and G. Magnino. 1996. Effects of exogenous cAMP on the morphology and behavior of dissociated cells of the sponge Clathrina cerebrum (Porifera, Calcarea). Eur. J. Cell Biol. 70: 92–96.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

121

Garrone, R. and C. Lethias, Freeze fracture study of sponge cells. pp. 121–128. In: K. Rützler [ed.] 1990, New Perspectives in Sponge Biology. Smithsonian Institution Press, Washington. Garrone, R. and C. Lethias, and J. Escaig. 1980. Freeze-fracture study of sponge cell membranes and extracellular matrix. Preliminary results. Biol. Cell. 38: 71–74. Gauthier, M. and B. Degnan. 2008. The transcription factor NF-κB in the demosponge Amphimedon queenslandica : insights on the evolutionary origin of the Rel homology domain. Dev. Genes and Evol. 218: 23–32. Ghysen, A. 2003. The origin and evolution of the nervous system. Int. J. Dev. Biol. 47: 555– 62. Giovine, M. and M. Pozzolini, A. Favre, G. Bavestrello, C. Cerrano, F. Ottaviani, L. Chiarantini, A. Cerasi, M. Cangiotti, E. Zocchi, S. Scarfi, M. Sara, and U. Benatti. 2001. Heat stress-activated, calcium-dependent nitric oxide synthase in sponges. Nitric Oxide 5: 427–431. Green, C.R. and P.R. Bergquist. Cell membrane specializations in the Porifera. pp. 153–158. In: C. Lévi and N. Boury-Esnault [eds.] 1979. Biologie des Spongiaires. Editions du C.N.R.S., Paris. Green, C. and P.R. Bergquist. 1982. Phylogenetic relationships within the Invertebrata in relation to the structure of septate junctions and the development of occluding junctional types. J. Cell Sci. 53: 270–305. Grundfest, H. Evolution of conduction in the nervous system. pp. 43–86. In: A. Bass. [ed.] 1959. Evolution of Nervous Control from Primitive Organisms to Man. American Association for the Advancement of Science, Washington. Grundfest, H. Evolution of electrophysiological varieties among sensory receptor systems. pp. 107–138. In: J. Pringle. [ed.] 1965. Essays on Physiological Evolution. The Macmillan Company, New York. Hanström, B. 1928. Vergleichende Anatomie des Nervensystems der wirbellosen Tiere unter Berücksichtigung seiner Funktion. Springer, Berlin. Harrison, F.W. and L. De Vos. Porifera. pp. 29–90. In: F.W. Harrison and J.A. Westfall. [eds.] 1991. Microscopic Anatomy of Invertebrates. Wiley-Liss, New York. Hejnol, A. and R. Schnabel. 2006. What a couple of dimensions can do for you: Comparative developmental studies using 4D microscopy—examples from tardigrade development. Integr. Comp. Biol. 46: 151–161. Hernandez-Nicaise, M.-L. 1973. The nervous system of ctenophores III. Ultrastructure of synapses. J. Neurocytol. 2: 249–263. Hertwig, O. and R. Hertwig. 1878. Das Nervensystem und die Sinnesorgane der Medusen. Vogel, Leipzig. Holland, N.D. 2003. Early central nervous system evolution: an era of skin brains? Nat. Rev. Neurosci. 4: 617–627. Horridge, G. 1968. Interneurons: Their Origin, Action, Specificity, Growth, and Plasticity. WH Freeman, London. Ishida, N. and M. Kaneko, and R. Allada. 1999. Biological clocks. Proceedings of the Natl. Acad. of Sci. 96: 8819–8820. Jacklet, J.W. 1997. Nitric oxide signalling in invertebrates. Invert. Neurosci. 3: 1–14. Jacobs, D.K. and N. Nakanishi, D. Yuan, A. Camara, S.A. Nichols, and V. Hartenstein. 2007. Evolution of sensory structures in basal metazoa. Integr. Comp. Biol. 47: 712–723. Johnston, G. 1842. A History of British Sponges and Lithophytes. W.H. Lizard, Edinburgh. Jones, W.C. 1957. The contractility and healing behaviour of pieces of Leucosolenia complicata. Q. J. Microsc. Sci. 98: 203–217. Jones, C.W. 1962. Is there a nervous system in sponges? Biol. Rev. 37: 1–50. Kilian, E.F. 1967. Ortsveränderungen von Süßwasserschwämmen unter dem Einfluß von Licht. Verh. Dtsch. Zool. Ges. 31 (Suppl.): 395–401. Klotz, K.-N. 2000. Adenosine receptors and their ligands. Naunyn Schmiedebergs Arch. Pharmacol. 362: 382–391.

© 2011 by Taylor and Francis Group, LLC

122

Key Transitions in Animal Evolution

Larroux, C. and B. Fahey, D. Liubicich, V.F. Hinman, M. Gauthier, M. Gongora, K. Green, G. Worheide, S.P. Leys, and B.M. Degnan. 2006. Developmental expression of transcription factor genes in a demosponge: insights into the origin of metazoan multicellularity. Evol. & Devel. 8: 150–173. Lendenfeld, R.V. 1889. A Monograph of the Horny Sponges. Trübner and Co, Ludgate Hill, EC, London. Lentz, T.L. 1966. Histochemical localization of neurohumors in a sponge. J. Exp. Zool. 162: 171–180. Lentz, T.L. 1968. Primitive Nervous Systems. Yale University Press, New Haven. Lethias, C. and R. Garrone, and M. Mazzarana. 1983. Fine structure of sponge cell membranes: Comparative study with freeze-fracture and conventional thin section methods. Tissue & Cell 15: 523–536. Leys, S.P. and G.O. Mackie. 1997. Electrical recording from a glass sponge. Nature 387: 29–30. Leys, S.P. and G.O. Mackie. 1999. Propagated electrical impulses in a sponge. Memoirs of the Queensland Museum 4430: 342. Leys, S.P. and G.O. Mackie, and R.W. Meech. 1999. Impulse conduction in a sponge. J. Exp. Biol. 202: 1139–1150. Leys, S.P. and R.W. Meech. 2006. Physiology of coordination in sponges. Can. J. Zool. 84: 288–306. Leys, S.P. Sponge coordination, tissues, and the evolution of gastrulation. In: M. Custódio, G. Lôbo-Hajdu, E. Hajdu and G. Muricy. [eds.] 2007. Porifera Reserach: Biodiversity, Innovation and Sustainability. Série Livros 28. Museu Nacional, Rio de Janeiro. Leys, S.P. and G.O. Mackie, H.M. Reiswig, and W.S. David. 2007. The Biology of Glass Sponges. Advances in Marine Biology, Volume 52. Academic Press London, Oxford, Boston, New York and San Diego. Lieberkühn, N. 1859. Neue Beiträge zur Anatomie der Spongien. Arch. Anat. Physiol. 353– 358, 515–529. Lieberkühn, N. 1863. Über Bewegungserscheinungen bei den Schwämmen. Archiv für Anatomie, Physiologie und wissenschaftliche Medicin, 30: 717–730. Linder, J.U. and J.E. Schultz. 2002. Guanylyl cyclases in unicellular organisms. Mol. Cell. Biochem. 230: 149–158. Linné, C. von. 1767. Systema Naturae Editio duodecima, reformata. Insecta, Vermes. Holmiae, Amsterdam. Loewenstein, W.R. 1967. On the genesis of cellular communication. Dev. Biol. 15: 503–520. Lusche, D. and K. Bezares-Roder, K. Happle, and C. Schlatterer. 2005. cAMP controls cytosolic Ca2+ levels in Dictyostelium discoideum. BMC Cell Biology 6: 12. Mackie, G.O. 1970. Neuroid conduction and the evolution of conducting tissue. Q. Rev. Biol. 45: 319–332. Mackie, G.O. Is there a conduction system in sponges? pp. 145–152. In: C. Lévi and N. BouryEsnault. [eds.] 1979. Biologie des Spongiaires. Editions du C.N.R.S., Paris. Mackie, G.O. and I.D. Lawn, and M.P. De Ceccatty. 1983. Studies on hexactinellid sponges: 2. Excitability, conduction and coordination of responses in Rhabdocalyptus dawsoni. Philos. Trans. R. Soc. Lond. B 301: 401–418. Mackie, G.O. 1990. The elementary nervous system revisited. Am. Zoologist 30: 907–920. Marshall, W. Coelenterata, Porifera, Tetractinellidae; Tafel XLVII. In: R. Leuckart. [ed.] 1885. Zoologische Wandttafeln der wirbellosen Thiere. Th. Fischer, Kassel. Martindale, M.Q. and K. Pang, and J.R. Finnerty. 2004. Investigating the origins of triploblasty: `mesodermal‘ gene expression in a diploblastic animal, the sea anemone Nematostella vectensis (phylum, Cnidaria; class, Anthozoa). Development 131: 2463–2474. Matsuno, A. and H. Ishida, M. Kuroda, and Y. Masuda. 1988. Ultrastructures of contractile bundles in epithelial cells of the sponge. Zoolog. Sci. 5: 1212. Mattson, M.P. and A.J. Bruce-Keller. 1999. Compartmentalization of signaling in neurons: Evolution and deployment. J. Neurosci. Res. 58: 2–9.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

123

McNair, G.T. 1923. Motor reactions of the fresh-water sponge Ephydatia fluviatilis. Biol. Bull. 44: 153–166. Meech, R.W. and G.O. Mackie, Evolution of excitability in lower metazoans. In: G. North and J. Greenspann. [eds.] 2007. Invertebrate Neurobiology. Cold Spring Harbor Monograph Series 49. Cold Spring Harbor Laboratory Press, New York. Meech, R.W. 2008. Non-neural reflexes: sponges and the origins of behaviour. Curr. Biol. 18: R70–R72. Meinhardt, H. 2002. The radial-symmetric hydra and the evolution of the bilateral body plan: an old body became a young brain. BioEssays 24: 185–191. Minchin, E. Sponges. pp. 1–178. In: E.R. Lancester. [ed.] 1900. A Treatise on Zoology. Part II. The Porifera and Coelentera. Adam & Charles Black, London. Mitropolitanskaya, R. 1941. On the presence of acetylcholin and cholinesterases in the Protozoa, Spongia and Coelenterata. C. R. Acad. Sci. USSR 31: 717–718. Mori, M. 1994. Electron microscopic and new microscopic studies of hepatocyte cytoskeleton: Physiological and pathological relevance. J. Electron. Microsc. (Tokyo) 43: 347–355. Nguyen, L. and J.-M. Rigo, V. Rocher, S. Belachew, B. Malgrange, B. Rogister, P. Leprince, and G. Moonen. 2001. Neurotransmitters as early signals for central nervous system development. Cell Tissue Res. 305: 187–202. Nichols, S.A. and W. Dirks, J.S. Pearse, and N. King. 2006. Early evolution of animal cell signaling and adhesion genes. Proc. Natl. Acad. Sci. USA 103: 12451–12456. Nickel, M. 2001. Cell biology and biotechnology of marine invertebrates. Sponges (Porifera) as model organisms. Dissertation Thesis, Universität Stuttgart, Stuttgart. Nickel, M. and M. Vitello, and F. Brümmer. 2002. Dynamics of cellular movements in the locomotion of the sponge Tethya wilhelma. Integr. Comp. Biol. 42: 1285. Nickel, M. and F. Brümmer. 2002. Patterns of movement and their integration in sessile, nerveless metazoa illustrated by Tethya wilhelma (Porifera, Demospongiae). Zoology 105: 69. Nickel, M. 2004. Kinetics and rhythm of body contractions in the sponge Tethya wilhelma (Porifera : Demospongiae). J. Exp. Biol. 207: 4515–4524. Nickel, M. and F. Brümmer. 2004. Body extension types of Tethya wilhelma: cellular organisation and their function in movement. Boll. Mus. Ist. Biol. Univ. Genova 68: 483–489. Nickel, M. and J. Hammel, T. Donath, and F. Beckmann, Quantitative morphometrics and contraction analysis ot the marine sponge Tethya wilhelma—using synchrotron radiation based x-ray microtomography and in vivo x-ray imaging. pp. 1065–1066, Print/CDROM / Online: http://hasyweb.desy.de/science/annual_reports/2005_report/part1/ contrib/47/13996.pdf. In: U. Krell, J. Schneider and M. von Zimmermann [eds.] 2005, HASYLAB Annual Report 2005. Part I. DESY, Hamburg. Nickel, M. 2006. Like a ‘rolling stone’: quantitative analysis of the body movement and skeletal dynamics of the sponge Tethya wilhelma. J. Exp. Biol. 209: 2839–2846. Nickel, M. and E. Bullinger, and F. Beckmann. 2006a. Functional morphology of Tethya species (Porifera): 2. Three-dimensional morphometrics on spicules and skeleton superstructures of T. minuta. Zoomorphol. 125: 225–239. Nickel, M. and T. Donath, M. Schweikert, and F. Beckmann. 2006b. Functional morphology of Tethya species (Porifera): 1. quantitative 3D-analysis of T. wilhelma by synchrotron radiation based x-ray microtomography. Zoomorphol. 209–223. Nickel, M. and J. Hammel, T. Donath, and F. Beckmann. Elastic energy load into spicule reinforced extracellular matrix of the sponge Tethya wilhelma?—Evidence from synchrotron radiation based x-ray microtomography pp. 1279–1280, Print / CD-ROM / Online: http:// hasyweb.desy.de/science/annual_reports/2006_report/part1/contrib/47/17965.pdf. In: U. Krell, J. Schneider and M. von Zimmermann. [eds.] 2006c, HASYLAB Annual Report 2006. Part I. DESY, Hamburg. Nickel, M. 2007. Movements without muscles, information processing without nerves. JMBA GME 6: 8–9. Nickel, M. 2010. Evolutionary emergence of synaptic nervous systems: what can we learn from the non-synaptic, nerveless Porifera?, Invertebrate Biology, 129: 1–16.

© 2011 by Taylor and Francis Group, LLC

124

Key Transitions in Animal Evolution

Noll, F.C. 1881. Mein Seewasser-Zimmeraquarium. Zool. Garten (Frankfurt) 22: 168–177. Oberdisse, E. and E. Hackenthal, and K. Kuschinsky. 1997. Pharmakologie und Toxikologie. Springer-Verlag, Heidelberg. Pallas, P. 1766. Elenchus Zoophytorum sistens generum adumbrationes generaliores et specierum cognitarum succinctas descriptiones cum selectis auctorum synonymis. P. van Cleef, The Hague. Pallas, P. 1787. Charakteristik der Thierpflanzen. Raspesche Buchhandlung, Nürnberg. Pansini, M. and R. Pronzato. 1990. Observations on the dynamics of a Mediterranean sponge community. K. Ruetzler. [ed.] New Perspectives in Sponge Biology; Third International Conference on the Biology of Sponges, Woods Hole, Massachusetts, USA, November 17–23, 1985. ix+533 pp. 404–415. Pantin, C.F.A. 1952. The elementory nervous system. Proc. R. Soc. Lond. B. Biol. Sci. 140: 147–168. Pantin, C.F.A. 1956. The origin of the nervous system. Pubblicazione della Stazione Zoologica di Napoli 28: 171–181. Parker, G.H. 1910. The reactions of sponges, with a consideration of the origin of the nervous system. J. Exp. Zool. 8: 765–805. Parker, G.H. 1919. The Elementary Nervous System. Lippincot, Philadelphia. Passano, L. 1963. Primitive nervous systems. Proc. Natl. Acad. Sci. USA 50: 306–313. Pavans de Ceccatty, M. 1955. Le systeme nerveux des éponges calcaires et siliceuses. 288. Pavans de Ceccatty, M. 1959. Les structures cellulaires de type nerveux chez Hippospongia communis LMK. Ann. Sci. Nat. Zool. Biol. Anim. 12: 105–112. Pavans de Ceccatty, M. 1960. Les structures cellulaires de type nerveux et de type musculaire de l’éponge siliceuse Tethya lyncurium Lmck. C. R. Acad. Sci. Paris 251: 1818–1819. Pavans de Ceccatty, M. 1971. Effects of drugs and ions on a primitive system of spontaneous contractions in a sponge (Euspongia officinalis). Experientia 27: 57–59. Pavans de Ceccatty, M. 1974a. Coordination in sponges the foundations of integration. Am. Zoologist 14: 895–903. Pavans de Ceccatty, M. 1974b. The origin of the integrative systems a change in view derived from research on coelenterates and sponges. Perspect. Biol. Med. 17: 379–390. Pavans de Ceccatty, M. Cell correlations and integrations in sponges. pp. 123–135. In: C. Lévi and N. Boury-Esnault. [eds.] 1979. Biologie des Spongiaires. Editions du CNRS, Paris. Pavans de Ceccatty, M. 1981. Demonstration of actin filaments in sponge cells. Cell Biol. Int. 5: 945–952. Pavans de Ceccatty, M. 1986. Cytoskeletal organization and tissue patterns of epithelia in the sponge Ephydatia mülleri. J. Morphol. 189: 45–66. Pavans de Ceccatty, M. 1989. Les éponges, à l’aube des communications cellulaires. Pour la Science 142: 64–72. Perovic, S. and A. Krasko, I. Prokic, I.M. Mueller, and W.E.G. Mueller. 1999. Origin of neuronal-like receptors in Metazoa: Cloning of a metabotropic glutamate/GABA-like receptor from the marine sponge Geodia cydonium. Cell Tissue Res. 296: 395–404. Philippe, H., R. Derelle, P. Lopez, K. Pick, C. Borchiellini, N. Boury-Esnault, J. Vacelet, E. Renard, E. Houiliston, E. Quéinnec, C. Da Silva, P. Wincker, H. Le Guyader, S. Leys, D.J. Jackson, F. Schreiber, D. Erpenbeck, B. Morgenstern, G. Wörheide, and M. Manuel. 2009. Phylogenomics Revives Traditional Views on Deep Animal Relationships. Curr. Biol. 19: 1–7. Pick, K.S., H. Philippe, F. Schreiber, D. Erpenbeck, D.J. Jackson, P. Wrede, M. Wiens, A. Alie, B. Morgenstern, M. Manuel, and G. Worheide. 2010. Improved phylogenomic taxon sampling noticeably affects non-bilaterian relationships. Mol. Biol. Evol.: msq089. Pierobon, P. and R. Minei, P. Porcu, C. Sogliano, A. Tino, G. Marino, G. Biggio, and A. Concas. 2001. Putative glycine receptors in Hydra: a biochemical and behavioural study. Eur. J. Neurosci. 14: 1659–1666. Plinus. The natural history. In: J. Bostock and H.T. Riley [eds.] 1855. Taylor and Francis, London.

© 2011 by Taylor and Francis Group, LLC

The Pre-Nervous System and Beyond

125

Pronzato, R. 1999. Sponge-fishing, disease and farming in the Mediterranean Sea. Aquat. Conserv. 9: 485–493. Pronzato, R. 2004. A climber sponge. Boll. Mus. Ist. Biol. Univ. Genova 68: 549–552. Prosser, C.L. and T. Nagai, and R.A. Nystrom. 1962. Oscular contraction in sponges. Comp. Biochem. Physiol. 6: 69–74. Prosser, C.L. 1967. Ionic analyses and effects of ions on contractions of sponge tissues. Z. Vgl. Physiol. 54: 109–120. Ramoino, P. and P. Fronte, F. Beltrame, A. Diasporo, M. Fato, L. Raiteri, S. Stigliani, and C. Usai. 2003. Swimming behaviour regulation by GABAB receptors in Paramecium. Exp. Cell Res. 291: 398–405. Ramoino, P. and L. Gallus, S. Paluzzi, L. Raiteri, A. Diaspro, M. Fato, G. Bonanno, G. Tagliafierro, C. Ferretti, and R. Manconi. 2007. The GABAergic-like system in the marine demosponge Chondrilla nucula. Microsc. Res. Tech. 70: 944–951. Ramoino, P. and S. Scaglione, A. Diasporo, F. Beltrame, M. Fato, and C. Usai. 2004. GABAA receptor subunits identified in Paramecium by immunofluorescence confocal microscopy. FEMS Microbiol. Lett. 238: 449–453. Ramoino, P. and C. Usai, F. Beltrame, M. Fato, L. Gallus, G. Tagliafierro, R. Magrassi, and A. Diaspro. 2005. GABAB receptor intracellular trafficking after internalization in Paramecium. Microsc. Res. Tech. 68: 290–295. Ramon y Cayal, S. 1937. Recollections of my life. Memoirs of the American Philosophical Society 8: 1–638. Redi, C.A. and S. Garagna, M. Zuccotti, E. Capanna, and H. Zacharias. 2002. Visual Zoology. The Pavia collection of Leuckart’s zoological wall charts (1877). Ibis, Como. Reiswig, H.M. 1971. In-situ pumping activities of tropical Demospongiae. Mar. Biol. 9: 38– 50. Ruediger, T. and J. Bolz, Neurotransmitters and the development of neuronal circuits. In: D. Bagnard [ed.] 2007. Axon Growth and Guidance. Springer Science+Business Media, LLC, Landes Bioscience, New York. Ruppert, E. and R. Fox, and R. Barnes. 2004. Invertebrate Zoology (7. Auflage). Thomson Publishing, Brooks/Cole. Sakarya, O. and K.A. Armstrong, M. Adamska, M. Adamski, I.F. Wang, B. Tidor, B.M. Degnan, T.H. Oakley, and K.S. Kosik. 2007. A post-synaptic scaffold at the origin of the animal kingdom. PLoS ONE 2: e506. Sarà, M. and A. Sarà, M. Nickel, and F. Brümmer. 2001. Three new species of Tethya (Porifera: Demospongiae) from German aquaria. Stuttgarter Beitr. Naturk. Ser. A 631: 1–15. Schierwater, B. and R. DeSalle. 2007. Can we ever identify the Urmetazoan? Integr.Comp. Biol. 47: 670–676. Schmidt-Rhaesa, A. 2007. The Evolution of Organ Systems. Oxford University Press, Oxford. Schmidt, O. 1866. Zweites Supplement der Spongien des Adriatischen Meeres enthaltend die Vergleichung der Adriatischen und Britischen Spongiengattungen. Verlag von Wilhelm Engelmann, Leipzig. Schultz, J.E. and T. Pohl, and S. Klumpp, 1986. Voltage-gated Ca2+ entry into Paramecium linked to intraciliary increase in cyclic GMP. Nature 322: 271–273. Seipel, K. and V. Schmid. 2006. Mesodermal anatomies in cnidarian polyps and medusae. Int. J. Dev. Biol. 50: 589–99. Sherrington, C.S. 1897. Croonian Lecture: The Mammalian Spinal Cord as an Organ of Reflex Action. Proceedings of the Royal Society of London 61: 220–221. Shibata, S. 2004. Neural regulation of the hepatic Circadian rhythm. Anat. Rec. Part A 280A: 901–909. Simpson, T.L. and G.A. Rodan. 1976. Role of cyclic AMP in the release from dormancy of freshwater sponge gemmules. Dev. Biol. 49: 544–547. Simpson, T.L. 1984. The Cell Biology of Sponges. Springer-Verlag, New York.

© 2011 by Taylor and Francis Group, LLC

126

Key Transitions in Animal Evolution

Smith, C. 1996. Sherrington‘s legacy: evolution of the synapse concept, 1890s–1990s. J. Hist. Neurosci. 5: 43–55. Sollas, W.J. 1888. Report on the Tetractinellida collected by H.M.S. Challenger during the years 1873–1876. Report on the Scientific Results of the voyage of H.M.S. Challenger 25: 1–458. Spring, J. and N. Yanze, C. Jösch, A.M. Middel, B. Winninger, and V. Schmid. 2002. Conservation of Brachyury, Mef2, and Snail in the Myogenic Lineage of Jellyfish: A Connection to the Mesoderm of Bilateria. Dev. Biol. 244: 372–384. Thiney, Y. 1972. Morphologie et cytochimie ultrastructurale de l‘oscule d‘Hippospongia communis LMK et de sa régénération. PhD thesis. Université Claude Bernard (Lyon I): 1–63. Trueman, E.R. 1975. The Locomotion of Soft-bodied Animals. Edward Arnold Limited, London. Tuzet, O. and R. Loubatiäres, and M. Pavans de Ceccatty. 1952. Les cellulues nerveuses de l‘éponge Sycon raphanus O.S. C. R. Acad. Sci. Paris 234: 1394–1396. Vacelet, J. 1966. Les cellules contractiles de l’éponge cornée Verongia cavernicola Vacelet. C. R. Acad. Sci. Paris 263: 1330–1332. Van Soest, R.W.M. Porifera, Schwämme. In: W. Westeheide and R. Rieger. [eds.] 1996. Spezielle Zoologie, Teil 1: Einzeller und Wirbellose. Fischer, Stuttgart. Walker, R.J. and L. Holden-Dye. 1991. Evolutionary aspects of transmitter molecules, their receptors and channels. Parasitology 102: S7–S29. Walker, R.J. and H.L. Brooks, and L. Holden-Dye. 1996. Evolution and overview of classical transmitter molecules and their receptors. Parasitology 113: S3–S33. Weissenfels, N. 1989. Biologie und Mikroskopische Anatomie der Süßwasserschwämme (Spongillidae). Gustav Fischer Verlag, Stuttgart. Weissenfels, N. 1990. Condensation rhythm of fresh-water sponges (Spongillidae, Porifera). Eur. J. Cell Biol. 53: 373–383. Weyrer, S. and K. Rutzler, and R. Rieger. 1999a. Serotonin in Porifera? Evidence from developing Tedania ignis, the Caribbean fire sponge (Demospongiae). Memoirs of the Queensland Museum 44: 659–665. Weyrer, S. and K. Rützler, and R. Rieger. 1999b. Serotonin in Porifera? Evidence from developing Tedania ignis, the Caribbean fire sponge (Demospongiae). Memoirs of the Queensland Museum 44: 659–665. Wiens, M. and B. Diehl-Seifert, and W.E.G. Müller. 2001. Sponge Bcl-2 homologous protein (BHP2-GC) confers distinct stress resistance to human HEK-293 cells. Cell Death and Differentiation 8: 887–898. Wilson, H.V. 1910. A study of some epitheloid membranes in monaxonid sponges. J. Exp. Zool. 9: 536–571. Wintermann, G. 1951. Entwicklungsphysiologische Untersuchungen an Süßwasserschwämmen. Zool. Jahr. 71: 428–486. Zocchi, E. and A. Carpaneto, C. Cerrano, G. Bavestrello, M. Giovine, S. Bruzzone, L. Guida, L. Franco, and C. Usai. 2001. The temperature-signaling cascade in sponges involves a heat-gated cation channel, abscisic acid, and cyclic ADP-ribose. Proc. Natl. Acad. Sci. USA 98: 14859–14864. Zocchi, E. and G. Basile, C. Cerrano, G. Bavestrello, M. Giovine, S. Bruzzone, L. Guida, A. Carpaneto, R. Magrassi, and C. Usai. 2003. ABA- and cADPR-mediated effects on respiration and filtration downstream of the temperature-signaling cascade in sponges. J. Cell Sci. 116: 629–636.

© 2011 by Taylor and Francis Group, LLC

Chapter 6

A Key Innovation in Animal Evolution, the Emergence of Neurogenesis: Cellular and Molecular Cues from Cnidarian Nervous Systems Brigitte Galliot

Summary The emergence of neurogenesis led to the acquisition of an efficient neuromuscular transmission in eumetazoans, as shown by cnidarians that use evolutionarily-conserved neurophysiological principles to crumple, feed, swim. However, the cnidarian neuroanatomies are quite diverse and reconstructing the urcnidarian nervous system is not an easy task. Three types of characters shared by anthozoans and medusozoans appear plesiomorphic: (1) three cell types forming the cnidarian nervous system, neurosensory cells, ganglionic neurons and nematocytes (cnidocytes) that combine mechano-chemosensation and venom secretion; (2) a chemical conduction through nerve nets and nerve rings, those being considered as annular central nervous systems; (3) a larval apical sensory organ that initiates metamorphosis. Other characters receive a disputed origin: (1) the neural stem cell(s), that are multipotent interstitial stem cell in hydrozoans, but not identified in other classes; (2) the electrical conduction Department of Zoology and Animal Biology, Faculty of Science, University of Geneva, Sciences III, 30 quai Ernest Ansermet, CH-1211 Geneva 4, Switzerland. E-mail: [email protected]

© 2011 by Taylor and Francis Group, LLC

128

Key Transitions in Animal Evolution

through neurons and epithelial cells so far detected only in hydrozoans; (3) the embryonic origin of the nervous system; (4) the medusa sensory organs, ocelli or lens-eyes for light, statocysts for pressure, which are lacking in anthozoans. Interestingly numerous gene families that regulate neurogenesis in bilaterian are expressed during cnidarian neurogenesis, e.g. cnidarian eyes express Pax, Six and opsin, supporting a common origin for vi